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Table of Contents Volume 1: Reproduction and Development 1.1 Sex Determination and the Development of the Genital Disc, Pages 1-38, L. Sánchez, N. Gorfinkiel and I. Guerrero 1.2 Oogenesis, Pages 39-85, D. A. Dansereau, D. McKearin and P. Lasko 1.3 Vitellogenesis and Post-Vitellogenic Maturation of the Insect Ovarian Follicle, Pages 87-155, L. Swevers, A. S. Raikhel, T. W. Sappington, P. Shirk and K. Iatrou 1.4 Spermatogenesis, Pages 157-177, R. Renkawitz-Pohl, L. Hempel, M. Hollmann and M. A. Schäfer 1.5 Gonadal Glands and Their Gene Products, Pages 179212, M. F. Wolfner, Y. Heifetz and S. W. Applebaum 1.6 Molecular Genetics of Insect Fertilization, Pages 213236, B. Loppin and T. L. Karr 1.7 Dosage Compensation, Pages 237-245, J. C. Lucchesi 1.8 Insect Homeotic Complex Genes and Development, Lessons from Drosophila and Beyond, Pages 247-303, L. K. Robertson and J. W. Mahaffey 1.9 Drosophila Limb Development, Pages 305-347, U. Weihe, M. Milán and S. M. Cohen 1.10 Early Embryonic Development: Neurogenesis (CNS), Pages 343-378, L. Soustelle and A. Giangrande 1.11 Development of Insect Sensilla, Pages 379-419, V. Hartenstein 1.12 The Development of the Olfactory System, Pages 421-463, G. S. X. E. Jefferis and L. Luo



Volume 2: Development 2.1 Myogenesis and Muscle Development, Pages 1-43, S. M. Abmayr, L. Balagopalan, B. J. Galletta and S. -J. Hong 2.2 The Development of the Flight and Leg Muscle, Pages 45-84, J. O. Vigoreaux and D. M. Swank 2.3 Functional Development of the Neuromusculature, Pages 85-134, D. E. Featherstone and K. S. Broadie 2.4 Hormonal Control of the Form and Function of the Nervous System, Pages 135-163, J. W. Truman 2.5 Programmed Cell Death in Insect Neuromuscular Systems during Metamorphosis, Pages 165-198, S. E. Fahrbach, J. R. Nambu and L. M. Schwartz 2.6 Heart Development and Function, Pages 199-250, R. Bodmer, R. J. Wessells, E. C. Johnson and H. Dowse 2.7 Tracheal System Development and Morphogenesis, Pages 251-289, A. E. Uv and C. Samakovlis 2.8 Development of the Malpighian Tubules in Insects, Pages 291-314, B. Denholm and H. Skaer 2.9 Fat-Cell Development, Pages 315-345, D. K. Hoshizaki 2.10 Salivary Gland Development and Programmed Cell Death, Pages 347-368, D. J. Andrew and M. M. Myat 2.11 Silk Gland Development and Regulation of Silk Protein Genes, Pages 369-384, E. Julien, M. Coulon-Bublex, A. Garel, C. Royer, G. Chavancy, J. -C. Prudhomme and P. Couble



Volume 3: Endocrinology 3.1 Neuroendocrine Regulation of Insect Ecdysis, Pages 1-60, D. Zitnan and M. E. Adams 3.2 Prothoracicotropic Hormone, Pages 61-123, R. Rybczynski 3.3 Ecdysteroid Chemistry and Biochemistry, Pages 125195, R. Lafont, C. Dauphin–Villemant, J. T. Warren and H. Rees 3.4 Ecdysteroid Agonists and Antagonists, Pages 197242, L. Dinan and R. E. Hormann 3.5 The Ecdysteroid Receptor, Pages 243-285, V. C. Henrich 3.6 Evolution of Nuclear Hormone Receptors in Insects, Pages 287-318, V. Laudet and F. Bonneton 3.7 The Juvenile Hormones, Pages 319-408, W. G. Goodman and N. A. Granger 3.8 Feedback Regulation of Prothoracic Gland Activity, Pages 409-431, S. Sakurai 3.9 Hormonal Control of Reproductive Processes, Pages 433-491, A. S. Raikhel, M. R. Brown and X. Belles 3.10 Hormones Controlling Homeostasis in Insects, Pages 493-550, D. A. Schooley, F. M. Horodyski and G. M. Coast 3.11 Circadian Organization of the Endocrine System, Pages 551-614, X. Vafopoulou and C. G. H. Steel 3.12 Hormonal Control of Diapause, Pages 615-650, D. L. Denlinger, G. D. Yocum and J. P. Rinehart 3.13 Endocrine Control of Insect Polyphenism, Pages 651-703, K. Hartfelder and D. J. Emlen 3.14 Biochemistry and Molecular Biology of Pheromone Production, Pages 705-751, G. J. Blomquist, R. Jurenka, C. Schal and C. Tittiger 3.15 Molecular Basis of Pheromone Detection in Insects, Pages 753-803, R. G. Vogt 3.16 Endocrinology of Crustacea and Chelicerata, Pages 805-842, E. S. Chang and W. R. Kaufman



Volume 4: Biochemistry and Molecular Biology 4.1 Insect Cytochrome P450, Pages 1-77, R. Feyereisen 4.2 Cuticular Proteins, Pages 79-109, J. H. Willis, V. A. Iconomidou, R. F. Smith and S. J. Hamodrakas 4.3 Chitin Metabolism in Insects, Pages 111-144, K. J. Kramer and S. Muthukrishnan 4.4 Cuticular Sclerotization and Tanning, Pages 145-170, S. O. Andersen 4.5 Biochemistry of Digestion, Pages 171-224, W. R. Terra and C. Ferreira 4.6 Lipid Transport, Pages 225-246, D. J. Van der Horst and R. O. Ryan 4.7 Proteases, Pages 247-265, M. R. Kanost and T. E. Clarke 4.8 Lipocalins and Structurally Related Ligand-Binding Proteins, Pages 267-306, H. Kayser 4.9 Eicosanoids, Pages 307-339, D. W. Stanley 4.10 Ferritin, Pages 341-356, J. J. Winzerling and D. Q. D. Pham 4.11 Tick-Talk, the Cellular and Molecular Biology of Drosophila Circadian Rhythms, Pages 357-394, P. H. Taghert and Y. Lin 4.12 Insect Transposable Elements, Pages 395-436, Z. Tu 4.13 Transposable Elements for Insect Transformation, Pages 437-474, A. M. Handler and D. A. O'Brochta 4.14 Insect Cell Culture and Recombinant Protein Expression Systems, Pages 475-507, P. J. Farrell, L. Swevers and K. Iatrou



Volume 5: Pharmacology 5.1 Sodium Channels, Pages 1-24, D. M. Soderlund 5.2 The Insecticidal Macrocyclic Lactones, Pages 25-52, D.Rugg, S. D. Buckingham, D. B. Sattelle and R. K. Jansson 5.3 Neonicotinoid Insecticides, Pages 53-105, P. Jeschke and R. Nauen 5.4 GABA Receptors of Insects, Pages 107-142, S. D. Buckingham and D. B. Sattelle 5.5 Insect G Protein-Coupled Receptors: Recent Discoveries and Implications, Pages 143-171, Y. Park and M. E. Adams 5.6 Scorpion Venoms, Pages 173-220, E. Zlotkin 5.7 Spider Toxins and their Potential for Insect Control, Pages 221-238, F. Maggio, B. L. Sollod, H. W. Tedford and G. F. King 5.8 Insecticidal Toxins from Photorhabdus and Xenorhabdus, Pages 239-253, R. H. ffrench-Constant, N. Waterfield and P. Daborn 5.9 Amino Acid and Neurotransmitter Transporters, Pages 255-307, D. Y. Boudko, B. C. Donly, B. R. Stevens and W. R. Harvey 5.10 Biochemical Genetics and Genomics of Insect Esterases, Pages 309-381, J. G. Oakeshott, C. Claudianos, P. M. Campbell, R. D. Newcomb and R. J. Russell 5.11 Glutathione Transferases, Pages 383-402, H. Ranson and J. Hemingway 5.12 Insect Transformation for Use in Control, Pags 403-410, P. W. Atkinson, D. A. O'Brochta and A. S. Robinson



Volume 6: Control 6.1 Pyrethroids, Pages 1-29, B. P. S. Khambay and P. J. Jewess 6.2 Indoxacarb and the Sodium Channel Blocker Insecticides: Chemistry, Physiology, and Biology in Insects, Pages 31-53, K. D. Wing, J. T. Andaloro, S. F. McCann and V. L. Salgado 

6.3 Insect Growth- and Development-Disrupting Insecticides, Pages 55-115, T. S. Dhadialla, A. Retnakaran and G. Smagghe 6.4 Azadirachtin, a Natural Product in Insect Control, Pages 117-135, A. J. Mordue (Luntz), E. D. Morgan and A. J. Nisbet 6.5 The Spinosyns: Chemistry, Biochemistry, Mode of Action, and Resistance, Pages 137-173, V. L. Salgado and T. C. Sparks 6.6 Bacillus thuringiensis: Mechanisms and Use, Pages 175-205, A. Bravo, M. Soberón and S. S. Gill 6.7 Mosquitocidal Bacillus sphaericus: Toxins, Genetics, Mode of Action, Use, and Resistance Mechanisms, Pages 207-231, J. -F. Charles, I. Darboux, D. Pauron and C. Nielsen-Leroux 6.8 Baculoviruses: Biology, Biochemistry, and Molecular Biology, Pages 233-270, B. C. Bonning 6.9 Genetically Modified Baculoviruses for Pest Insect Control, Pages 271-322, S. G. Kamita, K. -D. Kang, B. D. Hammock and A. B. Inceoglu 6.10 The Biology and Genomics of Polydnaviruses, Pages 323-360, B. A. Webb and M. R. Strand 6.11 Entomopathogenic Fungi and their Role in Regulation of Insect Populations, Pages 361-406, M. S. Goettel, J. Eilenberg and T. Glare 6.12 Pheromones: Function and Use in Insect Control, Pages 407-459, T. C. Baker and J. J. Heath

Volume : Indexes

EDITOR-IN-CHIEF LAWRENCE I. GILBERT University of North Carolina Chapel Hill, NC USA

EDITORS KOSTAS IATROU National Center for Scientific Research ‘‘Demokritos’’ Athens, Greece SARJEEET S. GILL University of California Riverside, CA USA

INTRODUCTION

The first edition of this treatise, entitled Comprehensive Insect Physiology, Biochemistry, and Pharmacology and edited by Gerald A. Kerkut (see p. ix for the Dedication to Prof. Kerkut ) and Lawrence I. Gilbert, was published in 1985. The preliminary meetings regarding that 13-volume series began in 1980 and it was agreed that there would be 12 volumes ‘‘that would provide an up-to-date summary and orientation on the physiology, biochemistry, pharmacology, behavior, and control of insects that would be of value to research workers, teachers, and students.’’ It was felt that the volumes would provide a classical background to the literature, include all the critical basic material, and then emphasize the literature from 1950 to the present day (1984). By mid-1981 most of the chapters were assigned to authors and the project was underway. It was agreed upon that there would be a final Volume 13, an Index volume, that combined the subject, species, and author indexes for all 12 volumes so that all material in those volumes could be located easily. This was a monumental undertaking and the 13 volumes finally appeared in 1985. It received plaudits from reviewers and from researchers around the world. One of us (LG.) was approached by Carrol Williams, an internationally renowned insect physiologist, at an International Congress of Entomology following the publication of the first edition, and Professor Williams noted that he had a choice between purchasing the 13-volume set for his laboratory and buying a new centrifuge. He then stated that he chose the 13-volume series and had never regretted it for a moment. There were more than 50 000 references to the literature in the first edition and more than 10 000 species of insects were referred to, and all of this information was readily available through the thirteenth volume (index). It is of interest that volumes of that original series are still being purchased some 19 years after this series was published. Over the past 5–6 years, one of us (LG.) was approached many times by colleagues around the world asking when a second edition of this series would be available. There was an obvious need because of the dramatic explosion of knowledge in insect science due to the utilization of molecular biological paradigms and techniques culminating in the elucidation of the Drosophila genome (fly database), the Anopholes genome, and of course this explosion continues. The use of this technology was really only beginning when the first series was developed and in 2000, 15 years after the publication of the first edition, we and Elsevier agreed that a new edition was necessary. This would be a series in which each chapter would begin by devoting a few pages summarizing the work in a particular research area up to the publication of the first series (1984–1985), and the remainder of the chapter would be a review of the literature from 1985 to 2003. It was agreed upon that each chapter would make full use of data arising from the utilization of molecular technologies if such data were applicable to that particular chapter subject. We chose to fulfill that premise in six volumes plus an index volume and that both a printed version and an electronic version would be available. As is evident, the title of the series has been changed, both to reflect the emphasis on molecular approaches and to alleviate the use of the obviously cumbersome title Comprehensive Insect Physiology, Biochemistry, Pharmacology, and Molecular Biology, which would have to be typed every time a researcher refers to a chapter in the series. We do not pretend, however, that this is a truly comprehensive treatise, since critical areas of research, e.g., ecology, evolutionary biology, and behavior, have not been included, but we have chosen to retain the use of the word ‘‘Comprehensive’’ for continuity with the first series.

viii

Introduction

The reader will note that the number of chapters in the six volumes is certainly well below the number of chapters in the original 12 volumes. The editors made very subjective decisions based on their own experience and knowledge regarding those areas where the most progress had been made over the past 18 years and omitted areas that have not grown significantly. Surely, there are areas of omission that many readers will feel were due to arbitrary decisions or lack of knowledge of the editors. That is bound to be the case although we spent a great deal of time deciding which chapters should be included, but there is little doubt that personal research bias played a role in those decisions. We are certainly confident that these volumes will be of great value to the research community, including postdoctoral investigators and graduate students. The new publishing technology will allow upgrades by the publisher every several years for those areas of research in which significant breakthroughs occur. This, of course, will only occur in the electronic format of the series and it is hoped that at least for the first few upgrades the original authors will take part. Lawrence I. Gilbert Kostas Iatrou Sarjeet Gill

DEDICATION

Professor Gerald Allan Kerkut: 19 August 1927–6 March 2004

Gerald Kerkut began his scientific career in Cambridge, graduating with a first class degree in Natural Sciences (1945–1948). He then elected to remain at Cambridge for his Ph.D. (1948–1951), studying locomotion in starfish under the supervision of Professor Eric Smith in the Department of Zoology. Gerald continued at Cambridge for a further three years (1951–1954) as a junior fellow of his college, Pembroke. During this period, he studied the electrical activity of the gastropod central nervous system, initially selecting slugs but, due to problems of identification, changed to the garden snail, Helix aspersa. A species he was to use for nearly 30 years. In 1954 Gerald moved to the University of Southampton to take up a lectureship in Animal Physiology within the Department of Zoology. Together with Professor Kenneth A. Munday, Gerald established the Department of Physiology and Biochemistry in 1959 and in 1966 was appointed to the second chair of Physiology and Biochemistry. He remained at Southampton throughout his academic life, retiring in 1992, when he was appointed Emeritus Professor. Gerald continued an active association with the University up until his death. He was invited by a number of universities to become chairman of their zoology or physiology departments but he always declined, preferring to remain at Southampton. Gerald Kerkut had a first class analytical mind, a prodigious capacity for hard work and an extensive knowledge of the scientific literature. He was always ready to challenge accepted dogma and this was very well illustrated in his inaugural lecture entitled ‘‘The Missing Pieces,’’ delivered in December 1968 (Kerkut, 1969). In this he reviewed his research on a number of topics, including the variable ionic composition of neurons, the role of the sodium–potassium pump in the maintenance of the resting potential, fast orthodromic and slow antidromic axon transport and amino acids as transmitters. Although much of his earlier research used the snail, Gerald also worked on insects where he was interested in the effect of sudden temperature changes on the nervous system. Using the cockroach leg preparation he observed that a sudden fall in temperature resulted in a transient increase in activity. Intracellular recordings showed that a decrease in temperature resulted in a depolarization of the membrane potential, which was responsible for the transient increase in activity. Q-10 values for changes in membrane potential of insect muscle were greater than those predicted from the Nernst equation. This led to his work on electrogenic metabolic pumps, summarized in his book with Barbara York, ‘‘The Electrogenic Sodium Pump’’ (Kerkut and York, 1971). Gerald also employed insect preparations for his research on glutamic acid as a transmitter and that certain insect neuron cell bodies possessed overshooting action potentials, for example, the octopamine-containing dorsal unpaired median (DUM) cells. Having been a great advocate of the use of isolated invertebrate central nervous system preparations, Gerald turned his attention to isolated mammalian preparations in the late 1970s. In particular, he developed the use of the isolated spinal cord in conjunction with Jeff Bagust. Gerald was particularly interested in sensory integration in the dorsal horn of the spinal cord. Although a great protagonist of

x Dedication

isolated preparations, Gerald stressed that observations using isolated preparations must always be related to the living animal. Gerald Kerkut was also very involved in the publication of scientific data both in terms of books and journals. In the late 1950s he met Robert Maxwell, the founder of Pergamon Press. They immediately formed an excellent rapport, and Maxwell encouraged Gerald to explore his idea of starting a journal in comparative physiology and biochemistry. To cut a long story short, this resulted in the publication of Comparative Biochemistry and Physiology with its first number in 1960 and Gerald as co-editor with Bradley T. Scheer. The journal proved a great success and eventually expanded into three sections, namely, Physiology, Biochemistry, and Pharmacology. Gerald continued to edit the journal until 1994. Another of his editing successes was Progress in Neurobiology, which he co-edited with John Phillis. Gerald also edited a very successful series of monographs in Zoology, which were published by Pergamon Press. Gerald’s interest in publishing work on insects is illustrated by his co-editing with Lawrence Gilbert in 1985 of the first edition of Comprehensive Insect Physiology, Biochemistry, and Pharmacology. He was particularly proud of the high standard of scholarship in these volumes. Gerald was always interested in evolution and with Maxwell’s support published ‘‘The Implications of Evolution’’ in 1960 (Kerkut, 1960). In this he examined the problem of the evolution and interrelationships of the animal phyla. This was a topic of lasting interest to him and one to which he returned in a mini-review entitled ‘‘Possible Evolutionary Futures for Mankind’’ (Kerkut, 1988). In 1988 a Festschrift was organized in Gerald’s honor and the proceedings published as a special edition of Comp. Biochem. Physiol. Gerald provided the opening chapter, in which he reviewed his research career up to that time (Kerkut, 1989). Gerald Kerkut enjoyed music, art, and travel. He was an accomplished pianist and had an extensive collection of art books. Up until 6–7 years ago, Gerald traveled widely in the Americas and the Far East. Gerald enjoyed teaching and interacting with undergraduates and postgraduates. Many will retain enduring memories of his humor and concerned interest in their welfare. During his active research period, Gerald trained over 80 postgraduates and their success will provide a lasting legacy to his memory.

References Kerkut, G.A., 1960. The Implications of Evolution. Oxford, Pergamon, p. 174. Kerkut, G.A., 1969. The Missing Pieces. University of Southampton, Southampton, UK, p. 15. Kerkut, G.A., 1988. Possible evolutionary futures for mankind. Comp. Biochem. Physiol. 90A, 5–10. Kerkut, G.A., 1989. Studying the isolated central nervous system; a report on 35 years: more inquisitive than aquisitive. Comp. Biochem. Physiol. 93A, 9–24. Kerkut, G.A., York, B., 1971. The Electrogenic Sodium Pump. Scientechnica, Bristol, UK, p. 182.

Robert Walker Head of Honours School School of Biological Sciences University of Southampton

PERMISSION ACKNOWLEDGMENTS

The following material is reproduced with kind permission of Nature Publishing Group Figure 11 of Chapter 1.8 Insect Homeotic Complex Genes and Development, Lessons from Drosophila and Beyond Figures 4 and 11 of Chapter 1.12 The Development of the Olfactory System Figures 3, 6, 7, 8 and 9 of Chapter 3.5 The Ecdysteroid Receptor Figure 5 of Chapter 3.11 Circadian Organization of the Endocrine System Figure 9 of Chapter 5.4 GABA Receptors of Insects http://www.nature.com/nature

The following material is reproduced with kind permission of American Association for the Advancement Figure 8 of Chapter 5.4 GABA Receptors of Insects http://www.sciencemag.org

Notes on the Subject Index To save space in the index the following abbreviations have been used: ETH – ecdysis triggering hormone GPCRs – G protein-coupled receptors PBAN – pheromone biosynthesis activating neuropeptide PDV – polydnaviruses PTTH – prothoracicotropic hormone QSAR – qualitative structure-activity relation RDL – resistance to dieldrin

1.1 Sex Determination and the Development of the Genital Disc L Sa´nchez, Centro de Investigaciones Biolo´gicas, Madrid, Spain N Gorfinkiel and I Guerrero, Universidad Autonoma de Madrid, Spain ß 2005, Elsevier BV. All Rights Reserved.

1.1.1. Sex Determination in Drosophila 1.1.1.1. Introduction and Historical Overview 1.1.1.2. The X/A Signal and the Activation of Gene Sex-lethal 1.1.1.3. The Sex Determination Genetic Cascade 1.1.1.4. Sex Determination Genes in Other Dipteran and Nondipteran Species 1.1.2. The Development of the Genital Disc of Drosophila 1.1.2.1. Historical Overview 1.1.2.2. Embryonic Organization of the Genital Disc 1.1.2.3. Organization and Patterning of the Genital Disc during Larval Development 1.1.2.4. Genetic Control of the Sexual Dimorphic Development of the Genital Disc 1.1.2.5. Gradual Acquisition of the Developmental Capacity to Differentiate Adult Structures 1.1.2.6. Evolutionary Considerations 1.1.3. Concluding Remarks and Perspectives

1.1.1. Sex Determination in Drosophila 1.1.1.1. Introduction and Historical Overview

Perpetuation by sexual reproduction is the rule within the animal kingdom. Males and females are different at the morphological, physiological, and behavioral levels. This sexual dimorphism results from the integration of the sex determination developmental program, which commits the embryo to either the male or the female pathway, and the developmental program which determines the type of cellular structure that will be differentiated in specific regions in the animal. This review deals with the genetic and molecular bases controlling sex determination and the sexual dimorphic development of the genital disc that gives rise to the terminalia, the most sexual dimorphic structure of the animal. It is focused on Drosophila since it is in this organism where a coherent picture about these developmental processes is now emerging. This has become possible thanks to the combination of sophisticated genetic and molecular biology techniques available in this model insect. However, sex determination and development of the genital disc in other insects will be also reviewed here. In Drosophila melanogaster, 2X;2A individuals (X, X chromosome; A, autosomal set) are females and XY;2A individuals (Y, Y chromosome) are males. Since females and males differ in the number of X chromosomes, a process has evolved to eliminate the

1 1 3 7 13 14 14 18 19 23 26 28 30

difference in the doses of the X-linked genes in the two sexes. This process is called dosage compensation (see Chapter 1.7). In D. melanogaster, the two X chromosomes in females are active and dosage compensation is achieved in males by hypertranscription of the single X chromosome. A series of results led to the discovery that in D. melanogaster sex is determined by the ratio of the X chromosomes to the sets of autosomes. First, XXY and X0 flies are female and male, respectively. This indicates that the Y chromosome plays no role in sex determination. Second, gynandromorphs are sexually mosaic individuals with some portions of the body typically male and others typically female. Such individuals arise from the loss of one X chromosome during the early development of XX flies. The sharp borderline between female and male tissues indicates that each individual cell chooses its sex autonomously, i.e., according to its chromosome constitution. Third, 2X;3A flies are mosaics with male and female structures. Collectively, these results indicate that gender in D. melanogaster is not determined by the absolute number of X chromosomes but by the ratio of X chromosomes to the sets of autosomes (X/A ratio signal). Supporting this contention, clones with one X chromosome and one set of autosomes (1X;1A clones) develop into female structures (Santamarı´a and Gans, 1980; Santamarı´a, 1983). In 2X;3A mosaic flies, the X/A ratio is at a threshold level between a normal female

2 Sex Determination and the Development of the Genital Disc

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and a normal male signal. Some cells interpret this ambiguous signal as female while others interpret it as male. Based on genotypes with variable X/A ratios, Bridges (1921, 1925) proposed his ‘‘balance concept’’ of sex determination. This hypothesis considers gender as a quantitative character, with continuous variation, under the control of two opposing polygenic signals, whose component factors have a small effect. The female-determining factors are located on the X chromosome and the male-determining factors are located on the autosomes. The opposing action of the two sets of signals would determine the gender of the fly, according to the stoichiometric assumption that two doses of the female-determining factors (two X chromosomes) outweigh the effect of two doses of male-determining factors (two sets of autosomes), leading to female development. However, two doses of male-determining factors (two sets of autosomes) outweigh the effect of one dose of femaledetermining factors (one X chromosome), leading to male development. In the case of 2X;3A flies, the stoichiometry of the interaction between femaleand male-determining factors is such that, within the same fly, in some cells the male development is imposed and in others the female factors prevail leading to female development. A breakthrough in the understanding of the genetic basis underlying sex determination in Drosophila was the work of Cline (1978), who found that Sex-lethal (Sxl) – a gene whose function depends on the X/A signal – is the key gene controlling both sex determination and dosage compensation processes, in such way that Sxl is activated in females (2X;2A) but not in males (X;2A). The function of Sxl would determine female sexual development and autosome-like basal transcription of each of the two X chromosomes in 2X;2A flies, whereas the lack of Sxl function would cause male sexual development and hypertranscription of the single male X chromosome (dosage compensation). Gadagkar et al. (1982) and Chandra (1985) proposed a model based on noncoding DNA to explain how cells would assess the X/A ratio signal. They proposed the existence of noncoding DNA sequences in the X chromosome (locus p) that would bind a repressor O encoded by an autosomal gene o. Moreover, they incorporated in their model Cline’s proposition about Sxl as the key gene controlling sex determination (Cline, 1978). In addition, they also incorporate Cline’s finding that the maternal product of gene daughterless (da) is also required for Sxl activation (Cline, 1978). The ‘‘noncoding DNA’’ model consists of the following components. First, the maternal Da product

that is stored in the egg; second, the O repressor that is encoded by the autosomal gene o and is activated in the zygote by the maternal Da product; and third, the gene Sxl and the locus p, which have affinity for the O repressor and RNA polymerase. A central assumption in this model is that the O repressor and RNA polymerase compete with each other for binding sites on both the Sxl and p loci in such a way that RNA polymerase binds to both loci with a lower affinity than that of the O repressor. It was further assumed that Sxl has a lower affinity than p for the repressor as well as the polymerase. According to this model, the amount of repressor in male and female embryos would be the same, since both have two copies of the gene o and both inherit the same amount of maternal Da product. The male embryo has only one X chromosome and therefore only one dose of the low affinity Sxl gene and one dose of the high-affinity locus p. The O repressor would bind significantly to both sites thus preventing the binding of RNA polymerase to Sxl. Consequently, little or no Sxl protein product would be produced. In the female embryo, there are two doses of both Sxl and p loci. Most of the O repressor will be sequestered by the p sites, so that the Sxl loci will be free to bind RNA polymerase. Synthesis of the Sxl product will then take place. The role of the X/A signal as the primary input for sex determination and dosage compensation could be visualized in two different ways. One possibility is that the X/A signal may be continuously needed during development for the cells to stay in the chosen sexual pathway and maintain the dosage compensation process properly adjusted. Under this hypothesis, X0 clones induced at any time during the development of XX flies would survive and differentiate male structures. Alternatively, the cells could use the X/A signal at a certain time in their development to set up their sex and dosage compensation processes. Under this second hypothesis, X0 clones induced before that time would survive and differentiate male structures, whereas X0 clones induced later would die because their dosage compensation process would be upset. To answer this question, a clonal analysis strategy was used. Genotypes were constructed that allowed the removal, by mitotic recombination induced by X-irradiation, of one of the two X chromosomes from an XX cell at different times in development. The results demonstrated that the X0 clones induced at around the blastoderm stage survive and differentiate male structures, while clones induced later in development die. However, the X0 clones survive and differentiate male structures if they carry a loss-of-function Sxl mutant allele, independent of

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Sex Determination and the Development of the Genital Disc

the time in development that the clones are induced. These results indicate that the X/A signal irreversibly sets, in a cell-autonomous manner, the state of activity of Sxl sometime around the blastoderm stage. Once this is achieved, the X/A signal is no longer needed and the activity of Sxl remains fixed (Sa´ nchez and No¨ thiger, 1983; Bachiller and Sa´ nchez, 1991). The capacity of the Sxl gene to function as a stable switch is due to a positive autoregulatory function of Sxl (Cline, 1984; Bell et al., 1991). Genetic evidence for this autoregulatory function came from the discovery of a new class of Sxl mutations that affected the sex determination process but still retained enough Sxl function to properly control dosage compensation. These Sxl mutant alleles showed the property that they do not need the maternal Da product for its expression in the zygote. In addition, they had the capacity of acting in trans causing the activation of a wild-type Sxlþ allele, which otherwise would not be activated because of the absence of maternal Da product (Cline, 1984). The posterior cloning and molecular characterization of Sxl demonstrated that this positive autoregulation is due to the requirement of the Sxl protein in the female-specific splicing of its own primary transcript (Bell et al., 1991). The gene Sxl controls the expression of two independent sets of regulatory genes (Lucchesi and Skripsky, 1981). The sex determination genes form one set, and mutations in these genes affect sex determination while having no effect on dosage compensation. The other set of genes is formed by the generically called male-specific lethal (msl) genes; mutations in these genes affect dosage compensation whilst having no effect on sex determination. This chapter is focused on sex determination in the somatic component of the animal (reviews: Cline and Meyer, 1996; Schu¨ tt and No¨ thiger, 2000). The germline exhibits sexual dimorphism, as does the somatic tissue. Cells with the 2X;2A chromosomal constitution will follow the oogenic pathway, while XY;2A cells will develop into sperm. Sex determination in the germline will not be reviewed here, and the interested reader is referred to the reviews of Cline and Meyer (1996) and Oliver (2002). Finally, Lucchesi reviews the dosage compensation process elsewhere in this Encyclopedia (see Chapter 1.7). 1.1.1.2. The X/A Signal and the Activation of Gene Sex-lethal

1.1.1.2.1. Genetic basis of the X/A signal The identification of a set of genes involved in the initial step of Sxl activation indicates that a conventional

3

genetic system is at the basis of the X/A signal. The isolation of X-linked genes involved in the formation of the X/A signal was approached by selection of sex-specific lethal mutations. This selection was based on the prediction that any relevant X-linked genes should function as numerator elements of the X/A signal. A numerator element should display several properties: 1. Reduction of its zygotic doses should specifically be lethal to 2X;2A flies (females) as a consequence of a failure to activate Sxl, which causes a hypertranscription of the two X chromosomes. This female-specific lethality should be suppressed by SxlM mutations. These are constitutive mutations that express female Sxl function independently of the X/A signal (Cline, 1978). 2. Increase of its zygotic doses should specifically be lethal to X;2A flies (males), because Sxl is inappropriately activated, leading to an alteration of dosage compensation. This male-specific lethality should be suppressed by loss-of-function Sxl mutations. 3. The activation of Sxl requires the maternal Da product (Cline, 1978). Therefore, mutations and any X-linked numerator of the X/A signal and a lower amount of maternal Da product are expected to display female-specific lethal synergistic interaction. Such interaction is also expected between mutations in different X-linked numerator genes of the X/A signal. In both cases, female lethality should be suppressed by SxlM mutations. 4. A variation of the zygotic doses of X-linked numerator genes should alter the sexual phenotype of triploid intersex (2X;3A) flies. An increase should feminize, whereas a reduction should masculinize these individuals. The expected properties for an autosomal denominator gene of the X/A signal are the opposite of those expected for an X-linked numerator gene. Genetic analyses have identified a set of zygotic and maternal genes that form the X/A signal. They fall into three classes. First, the X-linked numerator genes scute (sc, also called sisterless-b, sis-b) (Torres and Sa´nchez, 1989; Parkhurst et al., 1990; Erickson and Cline, 1991; Torres and Sa´nchez, 1991), sisterless-a (sis-a) (Cline, 1986), unpaired (upd, also called sisterless-c, sis-c) (Jinks et al., 2000; Sefton et al., 2000), and runt (run) (Duffy and Gergen, 1991; Torres and Sa´nchez, 1992). Second, the gene deadpan (dpn), the only autosomal denominator gene of the X/A signal that has been found so far (Younger-Shepherd et al., 1992). Finally, the maternal effect genes are daughterless (da) (Cline, 1978), hermaphrodite (her) (Pultz and

4 Sex Determination and the Development of the Genital Disc

Baker, 1995), extramacrochaetae (emc) (YoungerShepherd et al., 1992) and a gene(s) located in chromosomal region 7D10;7F12 of the chromosome (Sa´nchez et al., 1994). Not all of the genes involved in early Sxl activation play the same role. Among the X-linked numerator genes required for Sxl activation, only the sc and sis-a genes fulfil all criteria for numerator elements of the X/A signal. The autosomal gene dpn is the single known denominator element. The Xlinked genes run and upd also participate in the X/A signal, yet they do not play the predominant role played by sc and sis-a. Thus: 1. Simultaneous duplications of run and sc show very low Sxl-dependent male-specific lethality (Torres and Sa´ nchez, 1992) compared with the lethal effect produced by sis-a and sc duplications (Cline, 1988; Torres and Sa´nchez, 1989). 2. The gene run is only expressed in the trunk region of the embryo, whereas Sxl is activated in every cell of the embryo (Duffy and Gergen, 1991). 3. The majority of triploid intersexes heterozygous for run show normal male terminalia though some of them show a mixture of male and female structures (Duffy and Gergen, 1991; Torres and Sa´ nchez, 1992). In contrast, all triploid intersexes heterozygous for sc, for example, show completely normal male terminalia (Torres and Sa´ nchez, 1992). In the case of triploid intersexes heterozygous for sis-a, the degree of masculinization is almost complete (Cline, 1986). The terminalia are produced by the genital disc, and this is the most sexual dimorphic region of the fly (see below). 4. While mutations in sis-a and sc strongly downregulate SxlPe::lacZ expression, removal of upd activity or its downstream components has a significantly weaker effect, and residual SxlPe::lacZ expression is still seen in most mutant female embryos (Jinks et al., 2000; Sefton et al., 2000). 5. The effect of updþ transgenes is comparable to that of a chromosomal duplication of updþ, but is smaller than that seen for extra doses of sis-aþ or scþ. Eliminating upd activity reduces expression of SxlPe but to a lesser extent than eliminating sis-a or sc (Sefton et al., 2000). 6. The female-specific lethal synergistic effect between run and a deficiency of upd is weaker than the one between run and either sc or sis-a, or that between upd and either sc or sis-a (Sa´nchez et al., 1994, 1998). 7. Males doubly duplicated for a chromosomal region carrying upd and either sc or sis-a are as

viable as their brothers carrying only one of the duplications. This means that there is no malespecific lethal synergism between duplications at upd and either sc or sis-a (Sa´ nchez et al., 1994). 1.1.1.2.2. Molecular nature of the X/A signal The X/A signal acts on the early Sxl promoter and controls Sxl expression at the transcription level (Torres and Sa´nchez, 1991; Keyes et al., 1992). By definition, the X/A signal is strictly zygotic and the numerator X-linked products play the key role in forming this signal since the autosomal and maternal products are equally present in the male and female zygotes. Concerning the molecular nature of the X/A signal, only those products are considered for which a clear role has been found. The Sc (Villares and Cabrera, 1987; Murre et al., 1989), Da (Cronmiller et al., 1988; Murre et al., 1989), and Dpn (Bier et al., 1992) proteins contain a basic helix–loop–helix domain (bHLH). The HLH domain allows these proteins to form homodimers or heterodimers, while the basic domain is required for their binding to specific DNA sequences (Murre et al., 1989). Emc also belongs to the HLH family but is devoid of a basic domain (Ellis et al., 1990; Garrell and Modolell, 1990). SisA is a basic leucine-zipper (bZIP) protein (Erickson and Cline, 1993). Members of the bHLH and bZIP protein families are usually involved in transcriptional regulation (review: Massari and Murre, 2000). In vitro methods and yeast two-hybrid assays have allowed the analyses of protein–protein interactions. Experimental evidence supports the formation of the following homo- and heterodimers: Sc–Da (Cabrera and Alonso, 1991; Van Doren et al., 1991; Deshpande et al., 1995; Liu and Belote, 1995); SisA–Da (Liu and Belote, 1995); SisA–Dpn (Liu and Belote, 1995); Dpn–Dpn (Winston et al., 1999); Emc–Sc and Emc–Da (Van Doren et al., 1991). No interaction has been observed between Sc and Dpn (Deshpande et al., 1995; Liu and Belote, 1995) or between Sc and SisA (Liu and Belote, 1995). Nothing is known about the formation of higherorder multimers. 1.1.1.2.3. Role of the products that form the X/A signal The molecular characterization of the X/A signal components has clarified how they act on the early Sxl promoter (see Figure 1). The Sc and maternal Da products form complexes that induce early Sxl transcription by binding to a set of regulatory sites called E-boxes (Massari and Murre, 2000) within the early Sxl promoter (Estes et al., 1995; Hoshijima et al., 1995; Yang et al., 2001). The Emc protein forms heterodimer complexes

Sex Determination and the Development of the Genital Disc

5

Figure 1 Molecular basis of the X/A signal. The signal represents the ratio between activator and repressor complexes of the early Sxl promoter. For simplicity, it is assumed that in females (XX;AA), activators but not repressors are produced, whereas in males (X;AA), repressors but nor activators are formed, so that Sxl is only activated in females. Also, for the sake of clarity, representatives of each of the three classes of genes are chosen – X-linked zygotic, autosomal zygotic, and maternal genes – involved in early Sxl activation. X, X chromosome; III, third autosome; Y, Y chromosome. See text for explanation.

with either Sc or Da that are unable to bind to DNA (Davis et al., 1990; Voronova and Baltimore, 1990). Therefore, Emc prevents the formation of active Sc–Da complexes. The SisA protein is required for efficient induction of early Sxl transcription (Erickson and Cline, 1991; Estes et al., 1995) although specific binding sites for this protein on the early Sxl promoter have yet to be determined. Since the formation of SisA–Da (Liu and Belote, 1995) and SisA–Dpn (Liu and Belote, 1995) complexes has been reported, it is possible that SisA exerts a dual function: on one hand, it forms SisA–Da complexes that activate Sxl, and on the other, it sequesters Dpn through the formation of inactive SisA–Dpn complexes. Consequently, it lowers the amount of Dpn that remains free and capable of forming Dpn–Dpn dimers, which repress Sxl transcription by binding to a pair of adjacent regulatory sites in the early Sxl promoter (Estes et al., 1995; Hoshijima et al., 1995). 1.1.1.2.4. The organization of the early Sxl promoter The molecular organization of the early

Sxl promoter has been determined (Estes et al., 1995; Hoshijima et al., 1995; Yang et al., 2001). It has been shown that a fragment of 1400 bp upstream of the early Sxl transcription initiation site contains all the cis-acting sequences required for the control of early Sxl expression by the X/A signal. Deletion experiments suggest that the upstream distal region (1.4 to 0.8 kbp) is not essential for sex specificity. In contrast, the proximal region (390 bp to the transcription initiation site) is necessary and sufficient for transcriptional induction in presence of sufficient amounts of activators. The ostensible function of the distal region is to enhance transcription once it has been initiated by the proximal region; control of early Sxl transcription is mainly exerted through the proximal region (Yang et al., 2001). Binding sites controlled by the repressor have been termed D-boxes. Two D-boxes that bind Dpn dimers lie upstream and close to the transcription initiation site (Hoshijima et al., 1995; Yang et al., 2001). Due to their location close to the transcription start site, binding of the D-boxes is very

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6 Sex Determination and the Development of the Genital Disc

likely to inhibit the formation of the transcription machinery. 1.1.1.2.5. The action of the X/A signal on the early Sxl promoter The X/A ratio is primarily transduced by the relative amount of positive and negative regulators of Sxl (Parkhurst et al., 1990; Parkhurst and Ish-Horowicz, 1992). Although early Sxl protein has not been detected in males, suggesting that Sxl remains silent in males, it is still possible that Sxl transcription occurs in both sexes, but much more efficiently in females (activators outweigh repressors) than in males (repressors outweigh activators). As a result, early Sxl protein is abundantly produced in females whereas it remains undetectable in males. To explain in molecular terms how the X/A signal could exert its function, for simplicity it is considered that the X/A signal is such that, in females, activators but not repressors are formed whereas in males (X;2A), repressors but not activators are produced, so that early Sxl transcription occurs in females, whereas in males this gene remains silent (Figure 1). Since the interactions between some of the positive and negative regulators of Sxl have been shown, their relative abundance is regulated by a titration mechanism, where the existence of sequestering complexes like Sc–Emc and Da–Emc lowers the availability of free Sc and Da proteins for assembly into active complexes. Alternatively, the formation of SisA–Dpn would lower the amount of free Dpn and the formation of Dpn–Dpn repressors. Because the autosomal zygotic Dpn and the maternal Da and Emc are present in equal amounts in the two sexes, they have no discriminative power in sex determination. The sole difference between males (X;2A) and females (XX;2A) is the number of X chromosomes and, thus, the dosage of the X-linked gene products Sc and SisA. In females, the two doses of sc and sis-a produce sufficient Sc and SisA products to form Sc–Da and SisA–Da activator complexes, whereas negative Dpn–Dpn complexes are not formed due to the sequestration of Dpn by SisA into inactive SisA–Dpn complexes. Consequently Sxl is activated (Figure 1). In males, the amount of Sc and SisA products is half of that found in females. Emc sequesters Sc and Da, thus preventing formation of activator complexes. In addition, no SisA is available to sequester Dpn, so that the negative Dpn–Dpn complex is formed that prevents activation of Sxl. As mentioned above, other genes are also known to be required for early activation of Sxl: 1. Experimental evidence supports the idea that Run activates Sxl transcription by direct interaction

with the early Sxl promoter. First, Run-binding sites have been identified in the early Sxl promoter. Second, a mutant Run protein that cannot bind DNA fails to activate the early Sxl promoter. Finally, a modified Run protein containing a heterologous transcription activation domain activates rather than represses the early Sxl promoter, indicating that Run does not activate Sxl indirectly by repressing a Sxl-repressor. Nevertheless, it is not possible to exclude the possibility that the binding of Run to the early Sxl promoter interferes with the binding of a Sxl repressor (Kramer et al., 1999). 2. Both Upd and downstream components – the JAK kinase and the signal transducer and activator of transcription (STAT) – are needed for proper early activation of Sxl (Jinks et al., 2000; Sefton et al., 2000). Interestingly, the X/A signal acts cell-autonomously: the sexual pathway chosen by each cell depends on the cell’s chromosome constitution (X/A ratio). This is clearly observed in gynandromorphs (mosaic individuals with portions of the body typically male and others typically female) and in clones where the X/A ratio of the cells has been changed (see above). The question arises as to the function of the JAK/STAT signaling pathway (a noncellautonomous component) in a cell-autonomous decision process. It has been hypothesized that JAK/STAT signaling between adjacent cells may constitute a mechanism for stochastic compensation of differences in the activation of the early Sxl promoter by the X/A signal (Jinks et al., 2000). Indeed, gene transcription is a process of an intrinsically probabilistic nature (Zlokarnik et al., 1998; Fiering et al., 2000; Hume, 2000). 3. The maternal Her product is also necessary for early Sxl activation (Pultz and Baker, 1995), although it is not known whether specific binding sites for this protein exist in the early Sxl promoter or whether the protein forms complexes with any of the other positive regulators of Sxl. 1.1.1.2.6. Developmental meaning of the X/A signal The gene Sxl produces two temporallydistinct sets of transcripts corresponding to the function of its two promoters (Figure 2). The early set is produced as a response to the X/A signal, which controls Sxl expression at the level of transcription. The early promoter functions only around the blastoderm stage, when the X/A signal is formed. Later on, the late promoter starts functioning at the same level in both sexes and remains active throughout development and during adult life in both males and

Sex Determination and the Development of the Genital Disc

7

Figure 2 Scheme showing the two expression phases of Sxl related to its two promoters. PL and PE indicate the late and early promoters. E1 and L1–L5 indicate exons. The boxes and the thin lines represent exons (E1 and L1–L5) and introns, respectively. There are more exons (dotted line) that are included in the early and late Sxl mRNAs but these are not involved in the sex-specific and temporally specific splicing regulation of Sxl primary transcripts. Note that the early Sxl pre-mRNA follows a unique splicing pathway, whereas the late Sxl pre-mRNA follows two alternative splicing pathways.

females. In this case, the control of Sxl expression occurs by sex-specific splicing of its primary transcript. Male and female processed transcripts differ by the inclusion or exclusion of a male-specific exon (exon L3), which introduces a stop codon in the open reading frame, giving rise to a truncated, nonfunctional Sxl protein. In females, this exon is spliced out and normal, functional Sxl protein is produced (Bell et al., 1988; Bopp et al., 1991). Female-specific splicing of late Sxl pre-mRNA requires the presence of the functional Sxl product, and this constitutes the Sxl autoregulatory function (Bell et al., 1991). The early and late Sxl transcripts are distinct not only because of the different promoters used for their generation but, also, because of the different splicing pathways they follow (Figure 2). The early Sxl primary transcripts follow a fixed splicing pattern, where exon L2 and the male-specific exon L3 are excluded, and with the early specific exon E1 being directly spliced to exon L4. Exon L4 and the exons downstream from it are present in both early and late Sxl mRNAs. Splicing of the early Sxl primary transcript does not require the presence of the Sxl protein (Horabin and Schedl, 1996). The developmental meaning of the X/A signal is related to the existence of the two Sxl gene promoters. In females, early Sxl protein is abundantly produced, because the X/A signal (activators prevail over repressors) efficiently activates the early Sxl promoter (Figure 3). After the blastoderm stage, when the late Sxl promoter begins to function, the abundant early Sxl protein directs the first Sxl

pre-mRNA to the female-specific splicing pathway that gives rise to late functional Sxl protein, which subsequently ensures that the female-specific splicing state of Sxl is set up. In males, early Sxl protein is scarcely produced because the X/A signal (repressors prevail over activators) does not activate efficiently the early Sxl promoter. Consequently, when the late Sxl promoter starts functioning in males, the first Sxl pre-mRNA follows the male-specific splicing pathway that produces a truncated, nonfunctional Sxl protein. The result is the establishment of the male-splicing state of Sxl. Thus, the developmental meaning of the X/A signal is to provide the cells, at the beginning of development, with a sufficient quantity of early Sxl protein, which ascertains the female-splicing control of late Sxl pre-mRNA producing functional Sxl protein. 1.1.1.3. The Sex Determination Genetic Cascade

The epistatic relationships between the sex determination genes revealed that a hierarchical interaction exists among these genes (Baker and Ridge, 1980). The characterization of these genes showed that their expression during development is controlled by sex-specific splicing of their primary transcripts: the product of a gene controls the sex-specific splicing of the pre-mRNA of the downstream gene in the genetic cascade. Sxl is at the top of this cascade: its product controls the splicing of its own pre-mRNA and the splicing of the pre-mRNA from the downstream gene transformer (tra). The Tra product and the

8 Sex Determination and the Development of the Genital Disc

Figure 3 Scheme showing the developmental meaning of the X/A signal. For simplicity, it is assumed that the X/A signal is only produced in females. See text for details.

product of the constitutive gene transformer-2 (tra-2) control the sex-specific splicing of pre-mRNA from gene doublesex (dsx), which is transcribed in both sexes but gives rise to two different proteins, DsxF and DsxM, in females and males, respectively. The genes intersex (ix) and her are expressed in both sexes and in females, their products assist DsxF to activate female terminal differentiation and to repress male terminal differentiation, whereas Her helps DsxM to activate male terminal differentiation and to repress female terminal differentiation (Figure 4). The gene dsx is the last gene in the genetic cascade that controls the development of sexual somatic characters, yet there are exceptions. These concern the sex-specific characteristics of the central nervous system and the neuronal induction of a male-specific abdominal muscle, which depend on the function of tra and not dsx through the genes fruitless and dissatisfaction (Figure 4). The gene takeout, which is expressed in the fat body tissue, is closely associated with the adult brain function and is involved in male courtship behavior (Dauwalder et al., 2002). The determination of sexual behavior will not be reviewed here. The interested reader is referred to other reviews on the subject (Yamamoto et al., 1998; O’Kane and Asztalos, 1999; Billeter et al., 2002). 1.1.1.3.1. Control of Sxl expression by sex-specific splicing of its primary transcript The mechanism by which Sxl precisely controls the skipping of

its male-specific exon L3 is not totally understood. In contrast to most examples of exon-skipping events described in the literature, Sxl promotes a 100% switch between the two alternatively spliced forms of its own pre-mRNA, suggesting the existence of a complex mechanism of regulation. The role of the Sxl protein in the splicing of its own pre-mRNA is highlighted in brief. More extensive details have been reviewed in Penalva and Sa´ nchez (2003). The Sxl gene encodes one of the best-characterized members of the family of RNA-binding proteins. It has two segments, located in tandem in the central portion, with similarity to the RNA-binding domain sequences (RBD) or RNA recognition motifs (RRM). The amino terminus of Sxl is very rich in glycine. It has been shown that this domain is implicated in protein–protein interactions (Sxl multimerization) and absolutely required for proper control of Sxl RNA alternative splicing (Wang and Manley, 1997; Waterbury et al., 2000; Lallena et al., 2002). Deletions of the N- and C-termini does not interfere with the ability of the Sxl RBDs to properly bind in vitro to their target sequences, while both RNA-binding domains are required for sitespecific RNA binding (Wang and Bell, 1994; Kanaar et al., 1995; Sakashita and Sakamoto, 1996; Samuels et al., 1998). The Sxl-binding sequence consists of long stretches of poly(U) interrupted by two to four guanosines (Sosnowski et al., 1989; Inoue et al., 1990;

Sex Determination and the Development of the Genital Disc

9

Figure 4 Sex determination gene network. SxlF and SxlM stand for functional female and nonfunctional truncated male Sxl protein, respectively. TraF and TraM stand for functional female and nonfunctional truncated male Tra protein, respectively. DsxF and DsxM stand for functional Dsx protein in females and males, respectively. FruF and FruM stand for functional Fru protein in females and males, respectively. Dsf is dissatisfaction. To is takeout. In the absence of the X/A signal in males, the default state corresponds to the production of SxlM, TraM, and DsxM proteins. Ix and Her proteins are produced in both sexes. CNS, central nervous system. For description of the genes and their regulation see text.

Sakamoto et al., 1992; Horabin and Schedl, 1993a, 1993b; Sakashita and Sakamoto, 1994; Samuels et al., 1994; Singh et al., 1995; Wang and Manley, 1997). It has been observed that Sxl binds to RNAs containing two or more poly(U) sequences in a cooperative manner (Wang and Bell, 1994; Kanaar et al., 1995). Different models have been proposed for the molecular mechanism underlying the role of Sxl protein in controlling the female-specific splicing of its own pre-mRNA (Sakamoto et al., 1992; Horabin and Schedl, 1993a, 1993b; Wang and Bell, 1994; Wang and Manley, 1997; Waterbury et al., 2000; Penalva et al., 2001; Lallena et al., 2002; Nagengast et al., 2003). Nevertheless, there is a common aspect to all of them, namely, the fact that the Sxl protein participates in the skipping of the male-specific exon L3 (female-specific splicing of Sxl pre-mRNA) through binding to multiple Urich sequences placed in introns 2 and 3, surrounding the male-specific exon L3 (Figure 5). Multiple poly(U) sequences are also present in the adjacent introns of the male-specific exon L3 of the Sxl genes of D. virilis (Bopp et al., 1996) and D. subobscura (Penalva et al., 1996). The female-specific splicing of late Sxl pre-mRNA requires, in addition to the Sxl protein, the function of other genes, such as sans fille (snf ) (Albrecht and Salz, 1993; Flickinger and Salz, 1994; Salz and

Flickinger, 1996; Samuels et al., 1998), female-lethal-2-d (fl(2)d) (Granadino et al., 1990, 1992; Penalva et al., 2000; Ortega et al., 2003), and virilizer (vir) (Hilfiker and No¨ thiger, 1991; Hilfiker et al., 1995; Niessen et al., 2001). These genes, however, do not play any role in the splicing pattern of early Sxl transcripts (Horabin and Schedl, 1996). 1.1.1.3.2. Sxl protein as its own translation repressor Sxl gives rise to a variety of transcripts, some of them stage- and tissue-specific (Salz et al., 1989; Samuels et al., 1991; Keyes et al., 1992; Penalva et al., 1996). The Sxl mRNAs also contain several Sxl-binding sites at the 30 unstranslated region (UTR). It has been shown that the association of Sxl protein with the 30 UTR of Sxl mRNA significantly decreases Sxl protein expression (Yanowitz et al., 1999). Transcripts having a short 30 UTR are more abundant in the germline and during the early stages of embryogenesis (5–8 h postfertilization) (Salz et al., 1989). In later stages of development and during adult life, the most abundant transcripts have a long 30 UTR (up to 3600 nt) containing several Sxl binding sites (13 in the largest transcript) (Samuels et al., 1991). It was speculated that the negative autoregulatory loop of Sxl might play an important homeostatic role and that the presence of short transcripts at

10 Sex Determination and the Development of the Genital Disc

Figure 5 Sex-specific splicing of late Sxl pre-mRNA (autoregulatory Sxl function throughout development and adult life). Only the key exons (L2–L4) and introns (I2–I3) of the pre-mRNA are shown. The dot inside exon L3 represents translation stop codons. The hatched ellipsoids in introns I2 and I3 represent the sequences bound by the Sxl protein. Exons and introns are not drawn to scale and the numbers and distribution of Sxl-binding sequences are also not drawn to scale.

early stages of development allows a substantial amount of Sxl protein to be produced and a positive autoregulatory loop to be established via splicing. Once accumulation of the Sxl protein is no longer required, short mRNAs are substituted for larger ones whose expression can be repressed by Sxl. The balance between the negative and the positive posttranscriptional controls keeps the concentration of Sxl at levels that do not interfere with other cellular functions but are sufficient for regulation of Sxl splicing and splicing of its target genes (Yanowitz et al., 1999). Indeed, it has been shown that high levels of Sxl protein have toxic consequences for the cells (Meise et al., 1998; Saccone et al., 1998). 1.1.1.3.3. The role of transformer in sex determination The tra gene is transcribed in both sexes, but its RNA follows alternative splicing pathways (Figure 6). Intron 1 of tra has two alternative 30 splice sites. A non-sex-specific transcript is generated when the proximal 30 splice site is used. Use of this splice site introduces a stop codon in the open reading frame, leading to the production of a truncated, nonfunctional peptide. In females, approximately half of the tra pre-mRNA is spliced differently due to the intervention of the Sxl protein. In this case, the distal 30 splice site is used. As a result, the stretch containing the termination codon is

not included in the mature transcript and synthesis of full length Tra protein occurs (Boggs et al., 1987; Belote et al., 1989) (Figure 6). It has been determined that Sxl regulates femalespecific tra pre-mRNA splicing by a blockage mechanism rather than by enhancing the use of the female specific 30 splice site (distal 30 ss). There is a Sxl binding site at the polypyrimidine tract of the nonsex-specific splice site (proximal 30 ss) (Sosnowski et al., 1989; Inoue et al., 1990). This region is also the binding site for the U2 auxiliary factor (U2AF), an essential splicing factor important for the recognition of the 30 splice site. U2AF but not Sxl binds also to the polypyrimidine tract associated with the female-specific 30 splice site, but with 100 times less affinity (Valca´ rcel et al., 1993). Chimeric proteins containing the effector domain of U2AF fused to the complete RNA binding domain of Sxl inhibit rather than promote splicing to the non-sex-specific 30 splice site. This suggests that Sxl and U2AF compete for binding to the polypyrimidine tract associated with the non-sex-specific 30 splice site. Binding of Sxl to this sequence displaces U2AF, diverting it to the low affinity distal polypyrimidine tract and promoting the usage of the female-specific 30 splice site (Valca´ rcel et al., 1993; Granadino et al., 1997) (Figure 6). The chimeric U2AF–Sxl protein, however, does not disrupt Sxl pre-mRNA splicing regulation, in

Sex Determination and the Development of the Genital Disc

11

Figure 6 Sex-specific splicing of tra pre-mRNA. Boxes and thin lines represent exons (E1–E4) and introns, respectively. The dot inside exon E2 represents translation stop codons. The hatched ellipsoid in front of exon E2 represents the sequence to which both Sxl and U2AF proteins bind.

p0205

contrast to what occurs with tra splicing (Granadino et al., 1997). These data suggest that Sxl controls tra and Sxl pre-mRNA alternative splicing by different mechanisms. Notwithstanding, there is controversy concerning the function of the N-terminal region in tra RNA alternative splicing regulation. While it has been proposed that this region is not necessary for tra regulation (Granadino et al., 1997), others have proposed the opposite (Yanowitz et al., 1999). Differences in how the different groups have performed the experiments, which include different constructs (40aa deletion of the N-terminal region in one case and 94aa in the other), use of different promoters, and different real-time polymerase chain reaction (RT-PCR) methods to detect the splice forms, might explain the discrepancy. The genes female-lethal-2-d (fl(2)d) (Granadino et al., 1996) and virilizer (vir) (Hilfiker et al., 1995) are also required for female-specific splicing of the tra pre-mRNA. Genetic analyses have ruled out a direct role of sans-fille (snf ) in tra pre-mRNA splicing (Cline et al., 1999). 1.1.1.3.4. The role of doublesex, hermaphrodite, and intersex in sex determination At the bottom of the sex determination gene cascade are the genes dsx, her, and ix, which are expressed in both sexes.

The dsx pre-mRNA contains six exons: three common exons (E1–E3), a female-specific exon (E4), and two male-specific exons (E5–E6) (Figure 7). In females, the TraF protein, together with the constitutive Tra-2 protein (Goralski et al., 1989; Amrein et al., 1990; Mattox et al., 1990), directs the splicing of the dsx pre-mRNA to the female mode. The mRNA produced contains exons E1, E2, E3, and E4 and gives rise to the female DsxF protein, which promotes female sexual development. In males, where no functional Tra protein is available, the dsx pre-mRNA follows the male mode of splicing that gives rise to mRNA containing exons E1, E2, E3, E5, and E6, which produces male DsxM protein, which promotes male sexual development (Burtis and Baker, 1989; Hoshijima et al., 1991) (Figure 7). A further component that is crucial in the sexspecific splicing of dsx pre-mRNA, is the splicing enhancer dsxRE (doublesex repeat element) placed in the female-specific exon E4, 300 nt downstream of the 30 splice site (review: Maniatis, 1991). Six copies of a 13 nt repeat together with a purine-rich element (PRE) located between repeats 5 and 6, make up the dsxRE enhancer (Burtis and Baker, 1989; Lynch and Maniatis, 1996). The sequent of the female-specific 30 splice site in intron 3 departs significantly from the consensus 30

12 Sex Determination and the Development of the Genital Disc

Figure 7 Sex-specific splicing of dsx pre-mRNA. Boxes and thin lines represent exons (E1–E6) and introns, respectively. The dots inside exon E4 represent the dsxRE enhancer to which the TraF–Tra2 complex binds. Poly(A) indicates the polyadenylation site of the mRNA arising from female-specific splicing.

splice site and is weaker than the 30 splice site of the downstream intron 4. In males, the splicing machinery recognizes the 30 splice site of intron 4 instead of the 30 splice site of intron 3. Consequently, the male splicing follows and DsxM protein is produced. In females, however, because of the presence of TraF protein, TraF and Tra2, as well as the SR-protein RBP1, form a complex that binds to the dsxRE enhancer. This complex recruits U2AF and possibly other components of the general splicing machinery to the dsx pre-mRNA causing the splicing machinery to recognize the 30 splice site of intron 3 instead of the 30 splice site of intron 4. Consequently, female splicing follows and DsxF protein is produced (Hedley and Maniatis, 1991; Inoue et al., 1992; Tian and Maniatis, 1993, 1994; Zuo and Maniatis, 1996; Du et al., 1998; Nikolakaki et al., 2002). The activity and localization of the TraF, Tra-2, and RBD1 proteins depend on the activity of the LAMMER kinase encoded by the gene Darkener of apricot (Doa) (Du et al., 1998; Yun et al., 2000; Nikolakaki et al., 2002). The two Dsx proteins, DsxF and DsxM, are transcription factors controlling the activity of the final target genes involved in sexual differentiation. The two proteins share the N-terminal domain, which contains a DNA-binding domain (DM domain), whereas they differ at their C-terminal domains, which endow these proteins with their specific function (Burtis and Baker, 1989; Hoshijima et al., 1991). The her gene encodes a zinc finger protein, which endows to this protein the capacity to bind DNA, so

that Her can function as a transcription factor (Li and Baker, 1998a). This agrees with the dual function shown by the her gene. Its maternal expression is necessary for early Sxl activation (see Section 1.1.1.2.5), while its zygotic expression is necessary for female terminal differentiation and some aspects of male terminal differentiation (Pultz and Baker, 1995; Li and Baker, 1998b). The Dsx and Her products act either independently or interdependently to control different sexual differentiation processes. Thus, for example, the yolk protein (yp) genes yp1 and yp2 are the bestcharacterized female terminal differentiation genes. The expression of the yp1 and yp2 genes is under the control of gene dsx in the fat body (review: Bownes, 1994). DsxF and DsxM directly activate and inhibit the yp genes, respectively. They exert their function through the fat body specific enhancer (FBE). It has been found that in females, DsxF and Her independently activate the yp genes and that Her acts through sequences different than those of the FBE enhancer (Li and Baker, 1998b). In males, DsxM prevents the activation of yp genes by Her (Li and Baker, 1998b). However, the genes dsx and her also function interdependently to control the female differentiation of the sexual dimorphic foreleg bristles and the pigmentation of the 5th and 6th tergites (Li and Baker, 1998b). The ix gene is transcribed in both sexes and its pre-mRNA does not follow a sex-specific splicing, indicating that the Ix protein is present in both sexes. Ix shows similarity to proteins thought to

Sex Determination and the Development of the Genital Disc

act as transcriptional activators, although a DNAbinding domain has yet to be identified (GarrettEngele et al., 2002). By means of the yeast-2 hybrid and coimmunoprecipitation analyses, it has been possible to demonstrate that Ix interacts with DsxF but not DsxM. In addition, by gel shift analysis it has been shown that Ix and DsxF form a DNA-binding complex (Garrett-Engele et al., 2002). Ix also participates in the control of yp gene expression through the FBE enhancer (Garrett-Engele et al., 2002). Ectopic expression of DsxF in ix mutant males is not sufficient to induce expression of the yp genes, indicating that DsxF requires ix function to activate the yp genes (Waterbury et al., 1999). Collectively, these results suggest that Ix and DsxF form a complex to control female terminal differentiation. 1.1.1.4. Sex Determination Genes in Other Dipteran and Nondipteran Species

The search for genes homologous to sex determination genes of D. melanogaster has been undertaken. Among the genes that form the X/A signal, gene sc of D. subobscura (Botella et al., 1996) and gene sis-a of D. pseudoobscura and D. virilis (Erickson and Cline, 1998) have been characterized. These genes display a significant conservation in their structure and function. The Sxl gene has also been characterized in different Drosophila species. The structure and sequence organization of Sxl of D. virilis (Bopp et al., 1996) and D. subobscura (Penalva et al., 1996) have been determined. Like in D. melanogaster, Sxl regulation occurs by sex-specific alternative splicing: the Sxl transcripts in males have an additional exon containing stop translation codons. The Sxl of D. virilis, however, is unusual due to the presence in males of an open reading frame, downstream of the last stop codon in the male-specific exon, which encodes an Sxl protein. This is identical to the female Sxl protein except for the first 25 amino acids of the amino terminal region, which are encoded by differentially spliced exons. The male Sxl protein is predominantly accumulated in the embryonic ectoderm, suggestive of a putative role in the development of the central nervous system (Bopp et al., 1996). Sxl protein was also detected in males of other species (D. americana, D. flavomontana, and D. borealis) of the virilis radiation (Bopp et al., 1996). Outside the genus Drosophila, Sxl has been characterized in Chrysomya rufifacies (Mu¨ llerHoltkamp, 1995), Megaselia scalaris (Sievert et al., 1997, 2000), Musca domestica (Meise et al., 1998) and Ceratitis capitata (Saccone et al., 1998), which belong to the Brachycera Suborder, and in Sciara

13

ocellaris (Ruiz et al., 2003) which belongs to the Nematocera Suborder. Particularly interesting is the case of S. ocellaris, where as in D. melanogaster, gender depends on chromosome constitution: females are XX and males are X0 (Gerbi, 1986). Further, dosage compensation in Sciara appears to be achieved by hypertranscription of the single male X chromosome (da Cunha et al., 1994). The Sxl gene of all these species shows two main properties. First, Sxl is not regulated in a sex-specific fashion. Second, the Sxl proteins of the nondropsophilid and drosophilid species show a great degree of conservation in the two RBD domains and the few amino acids that separate them (the linker region). However, a high degree of conservation is not found outside these two domains. Collectively, these results suggest that the Sxl gene may not play the master regulatory role in sex determination in the nondrosophilids, as is the case with the Drosophila genus. This further suggests that Sxl was coopted to become the master regulatory gene in sex determination and dosage compensation during the evolution of the Drosophila lineage. The tra genes in the species that belong to the melanogaster group, D. simulans, D. mauritiana, D. sechellia, and D. erecta (O’Neil and Belote, 1992; Kulathinal et al., 2003), and D. hydei and D. virilis (O’Neil and Belote, 1992) have also been characterized. Its comparison with tra of D. melanogaster revealed an unusually high degree of divergence, yet the heterologous genes can rescue tra mutations in D. melanogaster. The gene tra-2 of D. virilis has been characterized as well (Chandler et al., 1997). It encodes a set of protein isoforms analogous to those of D. melanogaster, and it can rescue the tra-2 mutations in this species. Outside the genus Drosophila, tra has been characterized in C. capitata (Pane et al., 2002). Like in D. melanogaster, alternative splicing regulates the Ceratitis tra gene, so that only females contain a full-length protein. In contrast to Drosophila where Sxl regulates tra, however, the tra gene of Ceratitis shows an autoregulatory function that produces functional protein specifically in females. Therefore, tra in Ceratitis constitutes a cellular memory devise that maintains the female developmental pathway. The dsx genes of Megaselia scalaris (Sievert et al., 1997), Bactrocera tryoni (Shearman and Frommer, 1998), Bombyx mori (Ohbayashi et al., 2001; Suzuki et al., 2001), and Musca domestica (Hediger and Bopp, cited in Schu¨ tt and No¨ thiger, 2000) and C. capitata (Saccone et al., cited in Pane et al., 2002) have been characterized as well. In these species, dsx encodes male- and female-specific RNAs, which encode putative male- and female-specific Dsx

14 Sex Determination and the Development of the Genital Disc

proteins sharing the N-terminal region and differing at their C-terminal regions, like in Drosophila. Unlike the Drosophila case, however, the femalespecific intron in Bombyx dsx does not show a weak 30 splice site, and the Tra–Tra2 binding sequences (corresponding to the dsxRE enhancer in Drosophila; see Section 1.1.1.3.4) were not found. This suggests that the female-specific splicing of dsx pre-mRNA is the default splicing mode in Bombyx, in contrast to Drosophila, where the default splicing is male-specific. Collectively, these results agree with the model of Wilkins (1995), who proposed that during evolution, the sex-determining cascades were built from bottom to top, with the genes at the bottom being more conserved than the more upstream genes in the cascade.

1.1.2. The Development of the Genital Disc of Drosophila 1.1.2.1. Historical Overview

The imaginal discs of D. melanogaster are good experimental models for the study of developmental processes (No¨ thiger, 1972). Imaginal discs are formed by invagination from the embryonic ectoderm as pouches of cells and remain connected to the larval tissue until metamorphosis. They grow by cell division during the larval stages, whereby they acquire their patterning and typical shape. At the end of the larval period, each mature imaginal disc appears as an assembly of sets of spatially arranged cells with distinct determination states (patterning). These differentiate during metamorphosis as a response to hormonal stimulus (ecdysone). The result is the formation of the adult epidermal structures, such as the eye, antenna, leg, wing, and terminalia. Although these structures differ in appearance, the mechanisms controlling their growth and patterning, and even the genes involved, are very similar. The genital disc forms the terminalia, which comprise the entire set of internal and external genitalia and analia, the most sexually dimorphic structures in the adult (Figure 8). This disc shows certain characteristics that distinguish it from other imaginal discs. There are two imaginal discs, one for the right and one for the left, for each of the adult appendage except for the terminalia, which are derived from a single genital disc. The cells that form the genital disc are recruited from different segments in the embryo (see Section 1.1.2.3.1), whereas the other discs (with the exception of the eye–antenna disc) are formed by cells derived from single segments. Because of these characteristics, the

genital disc is well suited for studying developmental processes, such as the recruitment of cells to form complex structures and the control of patterning. Most importantly, however, the genital disc shows a unique feature. All imaginal discs except the genital one produce the same specific single adult appendage in both males and females. Thus, the eye, antenna, leg, wing, and haltere imaginal discs produce the corresponding adult structures in both sexes. In contrast, the genital disc has the potential to develop into either male or female terminalia, which are composed of different structures (Figure 8). Thus, the genital disc is a very good experimental model for the study of how sex determination genes, together with the homeotic ones – which determine cell identity – form an integrated genetic input that directs the development of a population of cells into two distinct, male or female, adult structures (reviews: Sa´ nchez and Guerrero, 2001; Christiansen et al., 2002). The complex organization of the genital disc was revealed through the analysis of gynandromorphs, flies composed of male (X0) and female (XX) cells. These animals allow the construction of fate maps concerning the relative position of the blastoderm founder cells for the different structures that compose the adult cuticle. The use of gynandromorphs to construct fate maps is based on an idea developed by Sturtevant (1929), who performed an analysis of gynandromorphs and calculated the frequencies with which two parts in the adult fly had different sexual phenotypes; i.e., whether they were formed by X0 or XX cells. The main conclusion was that the frequency with which the X0/XX border run between two parts was directly related to the distance of these parts in the adult and to the proximity of their founder blastoderm cells in the embryo (Janning, 1978). Dobzhansky (1931) and Kro¨ ger (1959) used gynandromorphs for the first time to determine the origin of the sexually dimorphic structures derived from the genital disc (Figure 8). These authors assumed that the male and female terminalia were homologous; i.e., the cells that composed the genital disc primordium have the capacity of developing along two alternative, male or female, pathways. Thus, cells with a X0 genetic constitution formed male terminalia, whereas cells having an XX constitution produced female terminalia. Under this assumption, it was expected to find a set of gynandromorphs that would cover a whole spectrum of terminal structures, ranging from pure male to pure female terminalia, through cases displaying different mixtures of pure male and female structures. The prerequiste to be fulfilled was that in the

Sex Determination and the Development of the Genital Disc

15

Figure 8 Scheme showing all the adult derivatives of the genital disc in both sexes ((a) female; (c) male) and photographs of external adult structures ((b) female; (d) male). External female terminalia: dAp, dorsal anal plate; vAp, ventral anal plate; T8, tergite eight; dVu, dorsal vulva; vVu, ventral vulva; dVp, dorsal vaginal plate; vVp, ventral vaginal plate. Internal female terminalia: U, uterus; Sr, seminal receptacle; Spt, spermatheca; Pov, parovaria; Od, oviduct (it is connected to the ovaries). Male external terminalia: Ap, anal plate; Ga, genital arch; Ll, lateral lobe; Lp, lateral plate; Cl, clasper; PA, penis apparatus; Ad, apodeme; Hy, hypandrium. Internal male terminalia: Ed, ejaculatory duct; Sp, sperm pump; Pg, paragonia (male accessory gland); Vdef, vas deferens (it is connected to the testes).

gynandromorphs the male and female structures should add up to a complete terminalia. However, Dobzhansly and Kro¨ ger noticed that this was not the case. No¨ thiger et al. (1977) addressed this question again more recently by analyzing the derivatives of mosaic genital discs of gynandromorphs. Their results constituted a benchmark in the comprehension of the organization and development of the genital disc. They found that, in the majority of the gynandromorphs, the male and female analia behave as expected if both were homologous and

derived from a common population of cells: the male and female anal structures added up to a complete analia. In contrast, the male and female genitalia yielded different results, as any female external genital structure could be combined with any male external structure. Furthermore, gynandromorphs were found that either lacked completely an external genital structure or had two external genitalia – a complete set of male and female structures. These observations applied not only to the external terminalia but also to the internal genital structures. The data were explained by the proposal

16 Sex Determination and the Development of the Genital Disc

that the genital disc is composed of three primordia: two separate primordia for the male and female genitalia, respectively, and a third one for the analia. The development of each genital primordium is either allowed or blocked depending on the chromosome constitution of their component cells. In X0 individuals, the male primordium develops to form the male genitalia and the development of the female primordium is blocked. In XX individuals, the female primordium develops to form the female genitalia, while the development of the male genital primordium is inhibited. However, the analia will always develop, giving rise to either male or female anal structures depending on the chromosome constitution of their cells, X0 or XX, respectively (but see below). According to this hypothesis the relative position of the three primordia that form the genital disc would be such that the female genital primordia occupies the most anterior position, the anal primordium occupies the most posterior position, and the male genital primordium is placed in the middle. This organization was suggested by the appearance of gynandromorphs lacking genitalia and having a female analia, and gynandromorphs that had complete female and male genitalia as well as male analia. In both classes of gynandromorphs, the X0/XX border runs between the female and the male genital primordia (Figure 9). In the first class, X0 cells populate the female genital primordium,

whereas XX cells populate the male genital and anal primordia. Consequently, the female and male genital primordia do not develop, whereas the anal primordium develops and produces a female analia. In the second class of gynandromorphs, the XX cells populate the female genital primordium whereas X0 cells populate the male genital and anal primordia. Consequently, the female genital primordium develops to produce complete female genitalia, whereas the male genital primordium develops and produces complete male genitalia, while the anal primordium develops and produces male analia. The hypothesized organization of the three primordia in the genital disc was confirmed when the disc’s embryonic origin was analyzed (Schu¨ pbach et al., 1978). It was found that the genital disc arises from a group of cells in the ventral side of the embryo. Within this group, three regions can be distinguished that are present in both sexes: an anterior region that gives rise to the female genitalia, a medial region that gives rise to the male genitalia, and a posterior one that produces the analia. Collectively, these data support the model that the genital disc is composed of three primordia: the female genital primordium (FGP) located anteriorly, the anal primordium (AP) located posteriorly, and the male genital primordium (MGP) placed in between (Figure 9). In each sex, only one of the two genital primordia develops depending on its sexual genotype, whereas the other remains in a so-called ‘‘repressed state,’’ and

Figure 9 Gynandromorphs lacking genitalia and having female analia (a), and gynandromorphs that have complete female and male genitalia plus male analia (b). In both classes of gynandromorphs, the X0/XX border runs between the female and the male genital primordia. FGP, female genital primordium; MGP, male genital primordium; AP, anal primordium. See text for explanation.

Sex Determination and the Development of the Genital Disc

does not form a recognizable adult structure (but see below). The ‘‘repressed’’ FGP in a male genital disc is termed RFP, and the ‘‘repressed’’ MGP in a female genital disc is termed RMP. In females, DsxF promotes the development of both the female genital and anal primordia, giving rise to the female genitalia and analia respectively; whereas the development of the male genital primordium is brought into the ‘‘repressed state’’ (RMP) (Figure 10). In normal males, DsxM promotes development of both the male genital and anal primordia, giving rise to the male genitalia and analia respectively, whereas the development of the female genital primordium is brought into the ‘‘repressed state’’ (RFP) (Figure 10). Mutations of the dsx gene, that do not produce DsxF and DsxM products, result in the generation of genital discs that show the same type of intersexual development in both 2X;2A and XY;2A flies. The latter is characterized by the development of both male and female genital primordia (though neither produce a complete set of genital structures) and by the anal primordium forming intersexual analia (Ehrensperger, 1983). The same intersexual development is observed when both DsxF and DsxM are simultaneously present within a cell (Epper, 1981) (Figure 10). This situation is encountered in 2X;2A flies of genotype dsxþ/dsxD, where the dsxþ allele produces the DsxF protein and the dsxD allele produces the DsxM protein (Nagoshi and Baker, 1990).

17

The demonstration that the RFP corresponds to the FGP (i.e., the localization of the RFP in the male genital disc) and that the RMP corresponds to the MGP (i.e., the localization of the RMP in the female genital disc) was performed by the analysis of intersexual genital discs and the clonal analysis of tra mutations. Thus: 1. The XX; dsxD/þ (Epper, 1981) and XX; dsx/dsx or XY; dsx/dsx or XX; ix/ix (Ehrensperger, 1983) intersexual genital discs contain three major regions (ventral, anterior dorsal, and posterior dorsal), which were isolated by fragmentation and transplanted into mature larvae ready to pupate, such that they underwent metamorphosis. The ventral fragment produced female genital structures, the dorsal anterior fragment produced male genital structures, and the dorsal posterior fragment produced anal structures. The ventral fragment of the intersexual genital disc corresponds to the region of the normal male genital disc that does not develop; i.e., to the RFP. The dorsal anterior fragment of the intersexual genital disc corresponds to the region of the normal female genital disc that does not develop; i.e., to the RMP. 2. The tra mutation transforms chromosomal (XX) females into males (see above). Clones homozygous for tra were induced by X-irradiation of

Figure 10 Sexually dimorphic development of the genital disc. A8, A9, and A10–11 indicate the corresponding abdominal segments. For further explanations on the role of Dsx, see text.

18 Sex Determination and the Development of the Genital Disc

XX; tra/þ larvae at late third larval instar. The genital discs were dissected and cultured in vivo in the abdomen of fertilized females to allow them to grow further before metamorphosis. Thereafter, the implants were dissected and, in a significant number of cases, the genital discs showed local overgrowths, at the site of the RMP in the female genital disc. These overgrowths were cut out of the discs and transplanted into larvae that underwent metamorphosis. All of them produced male genital structures. These results suggest that the RMP corresponds to the male genital primordium of the female genital disc that does not develop and that the tra function is needed to prevent development of the RMP (Epper and No¨ thiger, 1982). The cell lineage relationships between the primordia that compose the genital disc were studied by clonal analysis in the male and female genital discs. It was shown that clones overlapped in the right and left sides in both genital discs, but no clones were found that comprised genital and anal structures. This indicates that there is a cell lineage restriction between the analia and the genitalia in both sexes (Schu¨ pbach et al., 1978; Du¨ bendorfer and No¨ thiger, 1982; Janning et al., 1983). No cell lineage restriction between the two genital primordia could

be determined, since in each sex only one of the primordia develops (but see Section 1.1.1.2.2). 1.1.2.2. Embryonic Organization of the Genital Disc

As mentioned above, the embryonic region of the genital disc contains three different regions (Figure 11). The anterior region that contains precursors for the female genital primordium (FGP) corresponds to abdominal segment A8. The central region that contains precursors for the male genital primordium (MGP) corresponds to segment A9. And the most posterior region that gives rise to the analia of both males and females corresponds to segments A10 and A11 (Schu¨ pbach et al., 1978) (Figure 11). The embryonic genital disc can be distinguished at about 12 h of development by the expression of headcase (hdc) and escargot (esg), which mark all imaginal cells (Whiteley et al., 1992; Weaver and White, 1995; Casares et al., 1997). According to genetic mosaic analysis conclusions, the embryonic genital disc contains 12–15 cells (Hartenstein and Jan, 1992) that are organized as three clusters: two of them located to the left and right of the midline, and the other located posteriorly. These clusters do not correspond to the three primordia of the genital

Figure 11 Schematic representation of the compartmental organization of the female and male genital disc. Transverse sections (a, c) and dorsal and ventral view of (b, d) of the female (a, b) and male (c, d) genital disc respectively. Developing female genital primordium (FGP) or repressed female genital primordium (RFP) is in light gray; developing male genital primordium (MGP) or repressed male genital primordium (RMP) is in gray, and anal primordium (AP) is in dark gray. In each case, the anterior compartment (A) is lighter than the posterior compartment (P). The asterisk indicates that the P compartment cells of the three genital primordia are contiguous. For the rest of the symbols see legend to Figures 8 and 9.

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Sex Determination and the Development of the Genital Disc

disc; in fact, the tripartite organization of the genital disc is not obvious in the embryo (Casares et al., 1997). The genital disc grows during larval stages to give the sexually dimorphic disc characteristic of the third instar larvae. 1.1.2.3. Organization and Patterning of the Genital Disc during Larval Development

1.1.2.3.1. Segmental and compartmental organization of the genital disc As was previously described, early studies have demonstrated the existence of a cell lineage restriction between the analia and male or female genitalia (Schu¨ pbach et al., 1978; Du¨ bendorfer and No¨ thiger, 1982; Wieschaus and No¨ thiger, 1982). Initially, the question of a clonal restriction between male and female genitalia could not be addressed, because only one of the two genital primordia gives rise to adult structures in any one sex. Although it is now known that the RMP in females gives rise to the parovaria and that the RFP in males gives rise to a miniature 8th tergite (Keisman and Baker, 2001; see also below), these structures cannot be marked in the adult and are not suitable for analysis of clonal relationships. A cell lineage restriction analysis using dsx1 genital discs, where both primordia develop, has shown that clones could not cross the border between female and male genitalia (Gorfinkiel et al., 2003). Therefore, it was concluded that a cell lineage restriction exists between female and male genitalia, and between genitalia and analia. The evidence suggests that the three primordia of the genital disc represent three originally separate abdominal segments.

19

Each primordium of the genital disc has an anterior and a posterior compartment, which can be marked by the reciprocal expression of the cubitus interruptus (ci) and engrailed (en) genes (Freeland and Kuhn, 1996; Casares et al., 1997; Chen and Baker, 1997) (Figure 12). The ci and en mRNAs label the anterior and posterior compartments, respectively, and mark correspondingly the cells in all imaginal discs (Kornberg et al., 1985; Eaton and Kornberg, 1990). This agrees with en mutations affecting both male and female genitalia plus analia (Lawrence and Struhl, 1982; Epper and Sa´ nchez, 1983). The analysis of en loss-of-function clones has shown that the majority of external genital structures are of anterior origin (Casares et al., 1997; Emerald and Roy, 1998). Clones induced at the second larval stage do not cross between en mRNA- and ci mRNA-expressing cells (Chen and Baker, 1997) suggesting that at least by the beginning of the second instar, the genital disc has already subdivided into anterior and posterior compartments. A striking recent finding has challenged the idea that imaginal genital discs have, like all other imaginal discs, only anterior and posterior compartments. Ahmad and Baker (2002) have found a group of cells in the male genital primordium that express the breathless (btl) gene, which encodes the fibroblast growth factor receptor. These cells define a novel (third) compartment, because they do not express either ci or en and clones do not cross between btlexpressing cells and the rest of the disc. Surprisingly, these btl-expressing cells are not originally part of the male genital disc but are recruited into it during

Figure 12 Expression patterns of En, Dpp, and Wg in the female (a, b) and male (c, d) genital discs. The cells coexpressing Wg and Dpp are shown in black. (a) and (c) are ventral views, (b) and (d) are dorsal views of the corresponding genital discs. For the symbols see legend to Figures 9 and 11.

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20 Sex Determination and the Development of the Genital Disc

the larval stages. Cells in the male genital primordium expressing the gene branchless (bnl), which encodes the fibroblast growth factor, induce the btl-expressing mesodermal cells to migrate into the male disc. In this process, they lose the mesodermal marker twist (twi), acquire the ectodermal marker coracle (cora) and, eventually, form the paragonia. Therefore, three compartments compose the male genital primordium. Are the segmental borders different than the compartment borders? The genital disc is a good model to address this question, since the three primordia are contiguous to each other, like in the abdomen. It has been shown that in the wing disc, the Hedgehog (Hh) signaling pathway is responsible for the maintenance of the compartment border (see Chapter 1.9). smoothened (smo) anterior clones, which are unable to transduce the Hh signal, mix freely with posterior cells (Blair and Ralston, 1997; Rodriguez and Basler, 1997). In the genital disc, smo anterior clones can also mix with posterior cells but only from the same segment; smo anterior cells from one segment cannot mix with posterior cells from another segment (although they are contiguous). This lineage restriction does not depend on homeotic function either, although it cannot be ruled out that homeotic genes fix the restriction border much earlier during embryonic development (Gorfinkiel et al., 2003). Altogether, it can be concluded that segmental borders are different from the compartment borders. 1.1.2.3.2. Inductive signals in the development of the genital disc In the past decade, a paradigm has arisen about the genetic organization of imaginal discs. The selector gene en is expressed in posterior cells and activates the signal molecule Hh, which diffuses into the anterior compartment and elicits the activation of target genes. In the wing disc, Hh activates the expression of decapentaplegic (dpp), while in the leg and antennal disc, it activates de expression of dpp in dorsal anterior cells and wingless (wg) in ventral anterior cells. Both wg and dpp are signaling molecules that specify different identities in a concentration-dependent manner (reviews: Campbell and Tomlinson, 1995; Lawrence and Struhl, 1996) (see also Chapter 1.9). The same interactions take place in the genital disc, where each primordium is organized in a manner similar to that of the leg and antennal discs (Freeland and Kuhn, 1996; Casares et al., 1997; Chen and Baker, 1997; Sa´nchez et al., 1997; Gorfinkiel et al., 1999). In the genital and anal primordia, en in the posterior compartment activates the expression of hh, which, in turn, activates the expression of wg and dpp

in mutually exclusive and complementary domains, except in the A9 of female discs where dpp is not expressed (Freeland and Kuhn, 1996; Casares et al., 1997; Sa´nchez et al., 1997) (Figure 12). As it has been demonstrated in the leg disc (see Chapter 1.9), wg and dpp inhibit each other’s expression (Sa´nchez et al., 1997). The analysis of the expression domains of wg and dpp in the adult and the phenotypes obtained from lack of function and/or ectopic function of these signaling molecules have allowed the determination of the genital structures specified by these genes. Thus, in males, wg is expressed in the penis apparatus, the claspers, and the genital arch (Sa´ nchez et al., 1997). A requirement for wg in the development of these structures has been confirmed by the phenotype of wg hypomorphic alleles (Chen and Baker, 1997; Gorfinkiel et al., 2003). In females, the dorsal vaginal plates express high levels of wg and can be duplicated by ectopic expression of wg, suggesting that Wg specifies these structures (Chen and Baker, 1997; Sa´ nchez et al., 1997). The vulva has been found to require wg function also (Gorfinkiel et al., 2003). In addition, wg is required for the development of the internal genitalia in both sexes (Chen and Baker, 1997). The dpp gene is expressed also in the vaginal plates in females and in a subset of structures of the male genitalia like the genital arch, the hypandrium, and also the claspers (Sa´ nchez et al., 1997). The analysis of hypomorphic combinations of dpp has shown that dpp is required for all the genital structures except for the penis apparatus (Gorfinkiel et al., 2003). Furthermore, it has been shown that optomotor-blind (omb), a downstream gene in the dpp pathway (Grimm and Pflugfelder, 1996), is required for the formation of the dorsal bristles of the claspers and the hypandrium bristles (Gorfinkiel et al., 1999). It is also known that dachshund (dac), which is a downstream gene of wg in females but a downstream gene of dpp in males (Keisman and Baker, 2001; Sa´ nchez et al., 2001) (see Section 1.1.2.4.1), is required for proper formation of the internal ducts that connect the spermathecae to the uterus in females, and for correct formation of the claspers in males (Gorfinkiel et al., 1999; Keisman and Baker, 2001). The assignment of a particular signaling pathway to a specific adult structure is difficult because the Wg and Dpp pathways also act coordinately to activate downstream genes that, in turn, specify different structures. For example, the expression of Distal-less (Dll) is activated by the coordinated action of wg and dpp (Gorfinkiel et al., 1999), as happens in the leg disc (Dı´az-Benjumea et al., 1994)

Sex Determination and the Development of the Genital Disc

(see Chapter 1.9). Dll has been identified as a gene required for leg development and, in the terminalia, it is required for the development of the anal plates in both males and females (Sunkel and Whittle, 1987; Cohen and Ju¨ rgens, 1989; Gorfinkiel et al., 1997, 1999).

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f0065

1.1.2.3.3. Homeotic identity of the genital disc primordia The homeotic (Hox) genes specify structures along the anterior–posterior axis in most multicellular organisms (see Chapter 1.8). In Drosophila, the homeotic genes abdominal-A (abd-A), Abdominal-B (Abd-B), and caudal (cad) specify the identity of the three posterior primordia (Figure 13). The abd-A gene is expressed in part of the A8 primordium; in females this region correspond to the internal genitalia (Freeland and Kuhn, 1996). Based on its expression domain, it appears that abd-A is not required for the formation of the external genitalia (Sa´nchez-Herrero et al., 1985; Tiong et al., 1985). The Abd-B gene has two

21

transcripts: Abd-Bm and Abd-Br, which encode two proteins differing from each other in their Nterminal region. The m transcript is expressed in the A8 primordium, while the r transcript is expressed in the A9 primordium (Casares et al., 1997). The complete lack of Abd-B expression transforms the genitalia into leg or distal antenna, and this transformation is accompanied by ectopic expression of Dll and dachshund (dac), two genes required for leg development (Estrada and Sa´ nchez-Herrero, 2001) (see Chapters 1.8 and 1.9). Conversely, ectopic Abd-B expression transforms parts of the head into genitalia (Postlethwait et al., 1972; Kuhn et al., 1995). It has been suggested that the role of Abd-B is to specify genitalia by repressing Dll activity (Estrada and Sa´ nchez-Herrero, 2001). As Dll is activated by the combined action of wg and dpp in the genital disc, the role of Abd-B would be to counteract the activating function of these two signaling molecules. Because high levels of ectopic Dll are able to induce ventral appendages, legs, or antennae

Figure 13 Schematic representation of the control of dimorphic sexual development of male and female genital primordia by the dsx, Abd-B, and cad genes. A8, A9, and A10–11 indicate the 8th and 9th abdominal segments, and the fusion of 10th and 11th abdominal segments, respectively. The repression of Dpp expression in the RMP and the repression of the Wg pathway in the RFP are indicated by a continuous line. The activation of Wg and Dpp pathways are indicated by stippled lines in the FGP and MGP, respectively. See text for explanation. The broken line represents the anteroposterior compartment border in each primordia.

22 Sex Determination and the Development of the Genital Disc

(Gorfinkiel et al., 1997), the counteracting effect of Abd-B would be required to avoid leg development in the genitalia, and the Dll regulatory region would integrate the activating and repressing signals to control Dll transcription (Estrada and Sa´ nchezHerrero, 2001). The cad gene (Figure 13) is expressed in the anal primordium and gives rise to two different structures: the hindgut and the anal plates. This gene is required for the development of anal structures and, when ectopically expressed, can induce ectopic analia (Moreno and Morata, 1999). Because of this and despite the fact that cad is not physically linked to the Hox cluster, it is the fly’s most posterior Hox gene (Moreno and Morata, 1999). Consistently, cad clones show ectopic expression of Abd-B and concomitant transformation to genital structures in males (Moreno and Morata, 1999). Cad also activates Dll in the anal primordium; brachyenteron (byn), which is required for the development of the larval hindgut (Wu and Lengyel, 1998); and even-skipped (eve), another homeobox gene that is expressed in the presumptive region of the hindgut but is not required for hindgut development. Dll is also activated in the anal primordium by the combined action of Wg and Dpp, and is required for the formation of the anal plates (Gorfinkiel et al., 1999). Curiously, a mutually repressive interaction exists between Dll and eve in the anal primordium (Gorfinkiel et al., 1999). Thus, in the anal primordium, Cad acts in combination with Wg and Dpp to activate Dll (Moreno and Morata, 1999), while in the genital primordia, the Hox gene Abd-B counteracts the Wg and Dpp signals in order to downregulate Dll (Estrada and Sa´nchezHerrero, 2001) (see Section 1.1.2.3.4) (Figure 13). 1.1.2.3.4. Do the analia show a complex organization? Two chitinous plates, one dorsal and the other ventral to the anal opening, form the female analia. Two chitionous plates also form the male analia, one to the right and the other to the left of the anal opening (Figure 8). As mentioned above, the male and female analia behave as homologous structures derived from a common population of cells. Some experimental results, however, support the notion that the analia of both sexes may be derived from two distinct primordia, one giving rise to the dorsal analia in females or the complete analia in males (homologous primordium), while the other produces the ventral analia in females and no structure in males (nonhomologous primordium) (Andersen, 1979). A similar relationship has been described for the male and female analia in D. virilis (Newby, 1942).

First, XX;dsxD/þ, XX;dsx/dsx, or XX;ix/ix intersexes show one large horseshoe-shaped anal plate or separated left and right drop-shaped lobes, and a rudimentary plate with only two long bristles located ventral to the large plate (Andersen, 1979). Second, diplo-X homozygous tra-2ts animals reared at 21  C (intermediate temperature between femaleand male-determining temperature) develop a dorsal anal plate with the appearance of dorsally fused right and left male anal plates, while the ventral anal plate is greatly reduced or absent (Belote and Baker, 1982). Third, various degrees of sexual transformation in the analia were exhibited by diplo-X homozygous tra-2ts animals reared at temperatures intermediate between 16 and 29  C, or in animals that were shifted from 29 to 16  C at progressively later stages of larval development. With increasing degrees of masculinization (progressively later shifts), the ventral female anal plate becomes reduced, while the dorsal plate changes into the horseshoe-shaped form that was described above for intersexual flies, becoming split into a left and right plate in the most extreme transformation (Epper and Bryant, 1983). Fourth, the shift of XX; tra-2ts/tra-2ts animals from 16 to 29  C resulted in the quick formation of pure male anal plates. In the opposite shift, however, the formation of pure female anal plates occurred gradually. Moreover, the time course, required for the dorsal and ventral anal plates to show normal female phenotype, was different: when the dorsal anal plates were completely normal, it was still possible to find incomplete ventral anal plates (Sa´ nchez and Granadino, 1992). Finally, in flies mutant for Dll alleles with different degrees of functionality, the dorsal female anal plate and the two male anal plates are reduced, whereas the ventral female anal plate is normal. Furthermore, Dll clones do not develop anal plates in males or dorsal anal plates in females, but the development of the ventral female anal plates is not affected (Gorfinkiel et al., 1999). Thus, the dorsal female and male anal plates show the same genetic requirement for their development, supporting the idea that both are derived from a common primordium. It remains unknown, however, whether the segmental origin of this primordium is A10, and whether that of the ventral female analia forming primordium is A11. 1.1.2.3.5. Interaction between genital disc primordia The working model that emerges for the development of the genital disc is that each primordium develops as an independent unit in response to the general genetic cues that are also valid for other types of discs. However, there are also some results

Sex Determination and the Development of the Genital Disc

suggesting that the development of one primordium affects the development of the others. First, the regeneration of the genital disc, when different fragments of it are mixed (Littlefield and Bryant, 1979), suggests that some information is exchanged between primordia. Second, when cells of the anal or the genital primordia are killed by ectopic expression of the toxic ricin A gene, the development of the whole genital disc is impaired (Gorfinkiel et al., 2003). Similarly, mutations in Abd-Br, which is expressed only in A9 (see Section 1.1.2.4.4), produce a lack of female genitalia and analia (Casanova et al., 1986), and Abd-B mutations that transform the A8 and A9 segments into abdomen produce an absence of analia (Estrada and Sa´ nchez-Herrero, 2001). Third, because the genital disc results from the fusion of three adjacent segments, the posterior compartment of one segment is contiguous with the anterior compartment of the corresponding segment and the anterior compartment of the following one. Thus, it has been found that Hh in the posterior compartment diffuses in both directions and induces different target genes for and aft. While wg and/or dpp are induced towards the anterior, en expression is induced towards the following segment. These results suggest that signals can diffuse across the different primordia of the genital disc (Gorfinkiel et al., 2003). Accordingly, the ectopic expression of hh, wg, or dpp in the anal primordium induces duplications of genital structures. Also, dpp and wg mutant phenotypes that affect mainly the genitalia, can be rescued by expressing these genes in the analia. While the nonautonomous effect of Dpp is mediated by Dpp itself, the effect of Wg may be mediated by an as yet unidentified signal (Gorfinkiel et al., 2003). In summary, it can be concluded that the development of the different primordia of the genital disc is interdependent and that specific signals may diffuse across the segmental boundaries. 1.1.2.4. Genetic Control of the Sexual Dimorphic Development of the Genital Disc

1.1.2.4.1. Interaction of sex determination genes with signaling pathways Sexual differentiation deeply affects patterning and growth of the genital disc, suggesting that sex influences not only the terminal differentiation genes but also the early events in the morphogenesis of the genital structures. Accordingly, the development of the male and female genital disc requires the integration of the information supplied by the sex determination gene dsx, the morphogenetic signaling pathways, and the Hox genes. How this integration of patterning genes and Dsx takes place is starting to be elucidated.

23

The RMP of the DsxF-containing female genital disc does not express dpp (Freeland and Kuhn, 1996; Casares et al., 1997) (Figure 13). The lack of dpp expression in the RMP cannot be attributed to the failure of cells to receive the Hh signal, because Hh induces expression of wg in these cells. Rather, the dsx gene renders the cells of the male genital primordium capable or incapable of activating dpp. The cells of the male genital primordium are able to activate dpp in response to the Hh signal in male (absence of DsxF) but not female (presence of DsxF) genital discs (Figure 13). Thus, clones mutant for tra-2 (which is required for DsxF protein production) induced in the RMP of female genital discs, are always associated with dpp activation and show nonautonomous overgrowth (Sa´ nchez et al., 2001). Furthermore, it was also shown that the sex of the cells in the anterior compartment close to the anteroposterior compartment border (Hh responding cells), and not the sex of the whole genital disc, was the major factor controlling the growth of the genital primordia. When these cells are feminized in a male disc or masculinized in a female disc, both genital primordia respond by switching to growth patterns that reflect the sex in the induced anteroposterior compartment cells. Furthermore, these sexual changes in the Hh responding cells can change the expression of dpp in these cells (Keisman and Baker, 2001). Collectively, these results indicate that the developing or repressed status of the male genital primordium may be controlled by Dsx at the level of dpp expression, thereby dictating the sex specific patterns of growth within the genital disc (Keisman and Baker, 2001; Sa´ nchez et al., 2001). Moreover, it has been shown that DsxF is continuously required during female larval development to prevent activation of dpp in the RMP (Sa´ nchez et al., 2001) (see Section 1.1.2.4.2). The low proliferation rate of RFP in male genital discs cannot be attributed to the lack of Dpp or Wg, since both genes are expressed. Neither can it be due to a failure to respond to the Dpp signal, because the RFP expresses the Dpp-target gene omb (Gorfinkiel et al., 1999). Moreover, Dll, a Wg and Dpp target gene in the genital disc (Gorfinkiel et al., 1999), is not expressed in the RFP, where there is DsxM product, but is induced in the growing female genital primordium of genital discs that lack DsxM (Sa´ nchez et al., 2001). Thus, the repressed status of the female genital primordium seems to be controlled by blockage of the Wg signaling pathway by DsxM. Moreover, it has been also observed that DsxM is not continuously required during larval

24 Sex Determination and the Development of the Genital Disc

development for blocking the Wg-signaling pathway in the FGP (Sa´ nchez et al., 2001) (see Section 1.1.2.4.2). The sex determination genes also control the fibroblast growth factor (FGF) signaling cascade. The branchless (bnl) and the breathless (btl) genes, which encode FGF and its receptor, respectively, in Drosophila, are each expressed in two bilaterally symmetrical and adjacent groups of cells in the mature male but not female genital disc (Ahmad and Baker, 2002). The btl-expressing cells give rise to paragonia and vas deferens, which are major parts of the internal male genitalia (Figure 8). The induction of ectopic Traþ clones in the MGP of the male genital disc represses bnl expression. Conversely, in dsx mutant discs, where neither DsxF nor DsxM proteins are present, Bnl is expressed in the A8 and A9 segments. This indicates that DsxF represses the FGF pathway in the female genital disc and maybe DsxM activates this pathway in males (Ahmad and Baker, 2002). 1.1.2.4.2. The DsxM and DsxF products play positive and negative roles in the development of both the male and female genital primordia The established model for the development of the genital disc states that the male and female genital primordia, which are derived from two different cell populations, develop normally unless DsxF and DsxM, respectively, repress them (review: Steinmann-Zwicky et al., 1990). Accordingly, these two proteins play a negative, rather than positive, role in the developmental control of the genital disc. This model is primarily based on the observation that intersexual dsx mutant flies – simultaneously lacking DsxM and DsxF proteins – develop both male and female adult genital structures. These male and female genital structures are derived from the male and female genital primordia, respectively (recall that in these intersexual flies both the male and female genital primordia develop) (see Section 1.1.2.4.1). However, the inventory of these structures is not complete. This cannot be attributed to the mutual interference of male and female genital primordia, which develop simultaneously, because gynandromorphs (flies in which X0 male and XX female cells are observed within the same individual) with complete male and female adult genitalia have been found (No¨ thiger et al., 1977). Rather, it seems that in the dsx intersexual condition, both the male and female genital primordia fail to develop completely. Indeed, the number of cells in the mature dsx intersexual genital disc does not add up to the complete male plus female genital disc complement (Ehrensperger, 1983). This suggests that in the absence of DsxM

and DsxF products, both male and female genital primordia proliferate, albeit incompletely, and points to a positive role for DsxM and DsxF in the development of male and female genital primordia, respectively. Recent results concerning the expression of the gene dachshund (dac) in the genital disc (Sa´ nchez et al., 2001) support the standard model but also raise some uncertainties including one related to the positive role of the Dsx products in the development of the genital disc. In normal female genital discs, which have the DsxF product (Figure 13), dac is expressed in the Wg but not in the Dpp domain. In normal male genital discs, which have the DsxM product (Figure 13), dac is expressed in the Dpp but not in the Wg domain (Keisman and Baker, 2001; Sa´ nchez et al., 2001). In genital discs simultaneously lacking or having both DsxM and DsxF products, dac shows low levels of expression in both the Wg and Dpp domains. This agrees with the idea that dac is regulated by Wg in the female genital disc and by Dpp in the male genital disc, and that this differential regulation requires the function of dsx. This is also compatible with the idea that DsxM and DsxF have dual roles with respect to the expression of dac in the genital disc. In male genital discs, DsxM positively and negatively regulates dac expression in the Dpp and Wg domains, respectively. In female genital discs, DsxF positively and negatively regulates dac expression in the Wg and Dpp domains, respectively. At another level, Dsx has been also found to have an instructive and restrictive function. It was recently found that both the male and female primordia give rise to part of the adult structures in both sexes (Keisman and Baker, 2001). Using the expression patterns of genes such as en or wg as markers to track different genital primordia during metamorphosis, it was shown that each primordium gives rise to specific structures in each sex. In females, the male primordium forms a pair of female-specific accessory glands, whereas in males it produces a miniature eighth tergite. These findings indicate that, in one case, Dsx directs the female genital primordium to form either female genitalia or eighth tergite, and the male genital primordium to produce either male genitalia or the parovaria. Such a dual role of the Dsx products is not unique to the genital disc. A similar role has been observed in the regulation of the female-specific yolk protein (yp) genes: DsxM represses the yp genes in males, whereas DsxF activates the yp genes in females (see Chapters 1.2 and 1.3). In flies that lack both Dsx products, the yp genes are expressed at a basal level (Bownes and No¨ thiger, 1981; Coschigano and

Sex Determination and the Development of the Genital Disc

Wensink 1993; Waterbury et al., 1999). A positive role for DsxM has been also observed in the development of the sex comb in males (Jurnish and Burtis, 1993). 1.1.2.4.3. Different time requirements for the DsxM and DsxF products The following set of results suggests that the development of male and female genital primordia has different time requirements for DsxM and DsxF products: 1. Analysis of the growth of genital primordia and their capacity to differentiate adult structures in homozygous tra-2ts genital discs with two X chromosomes was performed using pulses of the male- and the female-determining temperatures in both directions during development. Regardless of the stage in development at which the female-determining temperature pulse was given (transient presence of functional Tra-2ts product, i.e., transient presence of DsxF product and absence of DsxM product), the male genital disc developed normal male adult genital structures and not female ones. This occurred even when the pulse was applied during pupation. Pulses at the male-determining temperature (temporal absence of functional Tra-2tts product, i.e., transient absence of DsxF product and presence of DsxM product), before the end of first larval stage produced female and not male genital structures. However, later pulses always gave rise to male genital structures, except when the pulses were given close to pupation. Further, the capacity of the female genital disc to differentiate adult genital structures was also reduced when the temperature pulse was applied during metamorphosis (Sa´ nchez et al., 2001). 2. When the effect of the male-determining temperature pulses was analyzed in the genital disc, it was found that overgrowth of the RMP was always associated with the activation of dpp in this primordium. However, this activation and the associated overgrowth only occurred when the temperature pulse was given after the end of first larval instar. This suggests that there is a time requirement for induction of dpp. The activation of this gene in the RMP and the cell proliferation that resumed in this primordium, as well as its capacity to differentiate into adult structures, was irreversible, since these were maintained upon return of the larvae to the female-determining temperature, when functional Tra-2ts product (i.e., presence of DsxF product and absence of DsxM product) was again available (Sa´ nchez et al., 2001).

25

3. The conclusion for the time requirement for induction of dpp was also supported by the fact that dsx11 clones (which lack DsxM) induced in the developing male genital primordium of XY; dsx11/þ male genital discs (which express only DsxM) after 24 h of development, differentiated normal male adult genital structures. However, when the dsx11 clones were induced in the time period between 0 and 24 h of development, they did not differentiate normally and gave rise to incomplete adult male genital structures. The different developmental capacity shown by the dsx11 clones depending on their induction time was explained as follows. When the clones were induced after 24 h of development, dpp was already activated. Indeed, these clones showed no change in the expression pattern of dpp or its targets (Sa´ nchez et al., 2001). Accordingly, these clones displayed normal proliferation and capacity to differentiate male adult genital structures. On the contrary, when the clones were induced early in development, dpp was not yet activated, as this gene is not expressed in the male genital primordium of male genital discs early in development (Casares et al., 1997). Therefore, when the male genital disc reaches the state in development when dpp is induced, the cells that form the clones activate this gene as in dsx mutant intersexual flies, because the clones have neither DsxM nor DsxF products. Consequently, these clones do not achieve a normal proliferation rate and do not produce normal adult male genital structures (Sa´ nchez et al., 2001). 4. The expression of the dac gene was analyzed in genital discs of diplo-X homozygous tra-2ts flies using pulses of the male- and the femaledetermining temperatures in both directions (recall that in male genital discs, DsxM positively and negatively regulates dac expression in the Dpp and Wg domains, respectively; and in female genital discs, DsxF positively and negatively regulates dac expression in the Wg and Dpp domains, respectively). It was found that the dac expression pattern switched from a ‘‘female type’’ to a ‘‘male type,’’ when male-determining temperature pulses were applied to tra-2ts larvae after the first larval instar. However, when the pulse was given during the first larval instar, dac was not activated in the Dpp domain of RMP, in spite of the fact that there was also a transient presence of DsxM instead of DsxF (Sa´ nchez et al., 2001). This was explained by the lack of competence of cells to express Dpp, which is acquired after the first larval instar (Casares et al., 1997).

26 Sex Determination and the Development of the Genital Disc

When the tra-2ts larvae reach such a developmental stage, these cells produce DsxF because they are returned to the female-determining temperature. DsxF prevents activation of dpp in the RMP and, consequently, no induction of dac expression occurs. In the female genital primordium, dac expression is strongly reduced in the Wg domain and is absent from the Dpp domain. Taken together, these results suggest that the development of male and female genital primordia have different time requirements for the DsxF and DsxM products. In females, DsxF is continuously required during female larval development for preventing activation of dpp in the RMP and, during pupation, for female cytodifferentiation. In males, DsxM seems not to be continuously required during larval development for blocking the Wg signaling pathway in the female genital primordium nor during pupation for male cytodifferentiation. 1.1.2.4.4. Integration of signaling pathways, sex determination, and homeotic information Homeotic and sex determination genes have to interact, along with the Hh, Dpp, and Wg signals, for normal development of the genitalia to occur (Keisman and Baker, 2001; Sa´ nchez et al., 2001). The dsx gene controls which of the two genital primordia will develop and which one will be repressed. Nevertheless, since DsxM and DsxF are equally active in each cell of the A8 and A9 segments of the male and female, respectively, another gene(s) is required to distinguish between female and male genitalia. The female genitalia develop from the A8 segment and the male from A9 (Freeland and Kuhn, 1996; Casares et al., 1997). Since the Abdominal-B (Abd-B) gene is responsible for the specification of these posterior segments (review: Duncan, 1996) (see Chapter 1.8), it has been proposed that AbdBr and Abd-Bm participate, together with dsx, in the sexually dimorphic development of the genital disc (Sa´ nchez et al., 1997, 2001). The genetic model that has been proposed for the control of the sexually dimorphic development of the genital disc (Figure 13), suggests that DsxM and DsxF combine with Abd-Bm and Abd-Br to make up the determining signals (Sa´nchez et al., 2001). First, it has been reported that Abd-B reduces the response to Wg and Dpp signals in the genital disc (Estrada and Sa´ nchez-Herrero, 2001). Abd-B seems to repress dac and Dll expression (perhaps by antagonizing the Dpp and Wg signaling) in some cells of the male and female primordia. It is still unknown if this multiple input acts directly on the dac and Dll regulatory regions. Second, in the

absence of both DsxM and DsxF products, there is basal expression of dpp and a basal functional level of the Wg signaling pathway in both the male and female genital primordia. This leads to both genital primordia developing, albeit incompletely. In normal females, the concerted input of DsxF and AbdBr causes repression of the development of the male genital primordium by preventing the expression of dpp and resulting in the RMP of female genital discs. In normal males, the concerted input of DsxM and Abd-Bm represses the female genital primordium by blocking the Wg signaling pathway, which gives rise to the RFP of the male genital discs. It is further proposed that DsxM plus Abd-Br increase dpp expression and the response to the Dpp signaling pathway in the MGP of male genital discs, and that DsxF plus Abd-Bm enhance the Wg signaling pathway function in the FGP of female genital discs. Therefore, DsxM plays a positive and a negative role in male and female genital primordia, respectively, while DsxF plays a positive and a negative role in female and male genital primordia, respectively. Such integration of the dsx and Abd-B functions seems not to be exclusive to the genital disc. The Drosophila 5th and 6th abdominal tergites are formed in both sexes, but in females they show dark pigmentation restricted to the posterior region (non-sex-specific pigmentation), whereas in males the pigmentation covers the whole tergite. The gene bric-a-brac (bab) is involved in the sexually dimorphic pigmentation of these tergites by integrating regulatory inputs from dsx and Abd-B (Kopp et al., 2000). Another integrated genetic input made up by dsx and the homeotic gene Sex combs reduced (Scr) has been invoked for sex-specific differentiation of the basitarsus of the prothoracic leg (see Chapter 1.9). This contains the sex comb, another malespecific structure (Jurnish and Burtis, 1993). However, it is not known at which level this interaction takes place. 1.1.2.5. Gradual Acquisition of the Developmental Capacity to Differentiate Adult Structures

At the end of the larval period, each mature imaginal disc represents a set of cells with different determination states that will be realized (differentiation) during metamorphosis as a response to the hormonal stimulus (ecdysone), resulting in the adult pattern that characterizes each imaginal disc (No¨ thiger, 1972). It has been reported, however, that imaginal discs acquire their capacity to differentiate into adult structures gradually throughout their larval growth; i.e., there is a specific temporal sequence in which the structures from imaginal discs

Sex Determination and the Development of the Genital Disc

of increasing age differentiate, this sequence being unique for each disc. These temporal sequences have been reported for the imaginal discs of the leg (Schubiger, 1974), the eye–antenna (Mindek and No¨ thiger, 1973; Gateff and Schneiderman, 1975), and the wing (Bownes and Roberts, 1979) (see Chapter 1.9). This was determined through transplantation of imaginal discs, from larvae at different stages of development, into mature larvae ready to pupate, and analyses of the inventories of adult structures derived from the implanted discs upon eclosion. The genital disc also shows a gradual acquisition of the developmental capacity to differentiate adult structures. The strategy used for the demonstration of the latter (Sa´ nchez and Granadino, 1992) involved a temperature-sensitive allele tra-2ts1 that was characterized because XX flies homozygous for this mutation develop as females at the permissive temperature (16  C), whereas they develop as males at the restrictive temperature (29  C) (Belote and Baker, 1982) (see Section 1.1.2.4.3 for the function of gene tra-2). The rationale of the strategy was as follows. When tra-2ts1 homozygous diplo-X larvae are raised at 16  C, when the Tra2 product is active, the FGP develops while the development of the MGP is blocked. If, during larval development, these larvae are shifted to 29  C, when no functional Tra-2 product will be available, the cells of the MGP will enter into a normal proliferation rate (Belote and Baker, 1982; Epper and Bryant, 1983). The degree of growth that the MGP reaches will depend on the time during development when the temperature shift is performed. Therefore, it is possible to establish a correlation between the development attained by the MGP and its capacity to differentiate adult structures. At the same time that the shift from 16 to 29  C prompts the MGP to develop, the cells of the FGP, which are developing at a normal rate, are brought into the slow proliferation rate that characterizes the RFP of the male genital disc, as a consequence of the inactivation of the Tra-2 product at the high temperature. Hence, it is possible to asses the developmental capacity attained by the FGP for differentiation of adult structures, when its normal growth is blocked at certain times before metamorphosis. An analogous situation occurs when the temperature shift goes from the 29 to 16  C but, in this case, the behavior of the female and male genital primordia is reversed. The results showed that the male genital primordium (male genital disc) follows a temporal sequence in its capacity to differentiate male external adult structures: the genital arch and lateral plates were the first structures that the male genital

27

primordium differentiated, followed by the clasper, hypandrium, and penis apparatus (see Figure 8 for the structures). Also, the female genital primordium (female genital disc) exhibits a sequence of capacity to differentiate external adult structures (8th tergite before vaginal plate), although in this case it is difficult to define a clear-cut sequence due to the reduced morphological elements that form the external adult female genital pattern. With respect to the analia, where cells proliferate independently of the state of tra-2 activity (Belote and Baker, 1982; Epper and Bryant, 1983), the temperature shift between 16 and 29  C, in both directions, indicated that the anal cells show a plasticity to change their sexual developmental program. Nevertheless, the change from female to male development occurred quickly, whereas that from male to female occurred gradually (Sa´ nchez and Granadino, 1992). There is a correlation between the sequence of structures differentiated by the genital disc and their topography in the fate map of the mature genital disc (Littlefield and Bryant, 1979; Ehrensperger, 1983; Epper and Bryant, 1983), as well as their cell lineage relationship (Du¨ bendorfer and No¨ thiger, 1982). It was suggested that this sequence is related to the dynamics of growth of the genital disc. It was further suggested that, at the end of the larval period, the mature genital disc is a mosaic of cells with different states of determination which, through metamorphosis, will differentiate into the distinct adult structures that form the terminalia. This mosaic pattern of the mature genital disc is not attained as an all-or-nothing event at the end of the larval period. Rather, during larval development, the proliferating genital disc is a mixture of cells already determined and cells that are still on their way to acquire a state of determination. The temporal sequence in the capacity to differentiate adult structures would reflect the temporal sequence of determination events in the genital primordia. In the temperature shift experiments of XX; tra2ts/tra-2ts animals, duplicated male genital structures corresponding to the penis apparatus were found. A similar duplication is observed in dsx or dsxD intersexual flies (Epper, 1981). What is the origin of these duplicated structures? The remaining vaginal plates always enclose the secondary set of male genitalia, whenever these female structures are formed. This location suggests that the duplicated male structures derive from the female genital primordium. Several results support this interpretation. Fate mapping of the dsxD intersexual genital discs showed that the small penis duplication maps on the female genital primordium (Epper, 1981). Moreover, Epper and Bryant (1983), in their experiments

28 Sex Determination and the Development of the Genital Disc

of culturing in vivo, at the male-determining temperature, fragments of XX; tra-2ts/tra-2ts genital discs that had been developing at the female-determining temperature, observed that the female genital fragment differentiated small penis structures. It is possible to argue that cutting errors during disc sectioning are the cause for the presence of the duplicated male structure. However, an independent observation supporting the notion that the penis duplication originates from the female genital primordium, stems from the analysis of the effect of en alleles in the genital disc (Epper and Sa´ nchez, 1983). The en1/en2 combination affects the female genital primordium, more severely, while the male genital primordium is most affected by the en2/en3 combination. In intersexual en2/en3; dsxD/þ flies, despite the lack of the entire penis apparatus in the male genital set, the small duplicated penis is always present. This rudimentary duplication, however, is absent from intersexual en1/en2; dsx/dsx flies, while the penis of the male genital set is always present. Thus, the secondary penis seems to be recognized by the en combinations as an element of the female genital primordium. Finally, in the above mentioned cases, the duplicated male structure is always accompanied by a reduction in the size of the ventral region of the vaginal plates. Collectively, these results suggest that a group of cells from the female genital primordium have the potential of following two alternative sexual pathways. In wildtype females, this group of cells gives rise to the ventral region of the vaginal plate, while in wildtype males it forms part of the ‘‘repressed’’ RMP primordium. In intersexual flies, when both female and male genital primordia develop, this group of cells produces the secondary penis. 1.1.2.6. Evolutionary Considerations

Evolutionary changes in morphology do not necessarily involve the generation of novel cell types but rather changes in morphogenesis, i.e., in the processes that are responsible for the generation of three-dimensional arrangements of various cell types (Raff and Kauffman, 1983). What is the genetic basis for morphological evolution? The genital disc, in addition to being a good experimental model in which to examine the control of pattern formation, is also well suited for the study of morphological evolution. In this respect, two aspects of the genital disc can be highlighted. First, the female external terminalia are very similar in the Drosophila species. The external male terminalia, however, display great differences; indeed, these are some of the main features used in taxonomy (Tsacas and Bocquet, 1976). Therefore, the terminalia are

particularly well suited for studying the great morphological variety observed in the male genitalia. The second aspect is related to the origin of the genital disc. 1.1.2.6.1. The genital disc in interspecific hybrids The genetic basis for the morphological evolution of the genital disc has been addressed through the analysis of interspecific hybrids of Drosophila. This analysis has focused on the effect of quantitative trait loci (QTL) on the shape of a particular structure of the male terminalia, the lateral lobe, in the sibling species D. melanogaster, D. simulans, D. mauritiana, D. sechellia, and their hybrids (Coyne, 1983; Coyne et al., 1991; Liu et al., 1996). The morphology of the lateral lobe differs markedly in these species. The genetic basis underlying the formation of this structure was studied through the production of first generation-interspecific hybrids. In the cases where the female hybrids were fertile, F2 backcrosses to males of either generation could be performed and it was possible to analyze hybrids that contained different combinations of genes of the two parental species. Evidence was presented supporting a polygenic basis for the shape of this male structure. Analysis of the morphological variation in other interspecific hybrids cannot be carried out because, in the majority of the cases, the species never interbreed, due to ethological isolation or other causes. A way of circumventing this problem is to crosstransplant germ cells. If the germ cells from the donor can develop in the soma of a host, it may be possible to obtain interspecific hybrids when the chimeric adult hosts are mated with individuals of their own species (Santamaria, 1977; Sa´ nchez and Schmid, 1984; Schmid et al., 1984; Lawrence et al., 1993). Following this strategy, germ cells from D. yakuba or D. teissieri – both belonging to the melanogaster subgroup (Lachaise et al., 1988) and not breeding with D. melanogaster – were transplanted into D. melanogaster host embryos, and the chimeric hosts were mated to D. melanogaster males. The germ cells of both donor species were capable of producing functional yakuba or teissieri oocytes, respectively, which, when fertilized by the melanogaster sperm, gave rise to yakuba–melanogaster and teissieri–melanogaster adult hybrids, whose external terminalia were characterized (Sa´ nchez and Santamaria, 1997). The female external terminalia are very similar in the three species, D. melanogaster, D. yakuba, and D. teissieri. The external male terminalia, however, display significant differences (Tsacas and Bocquet, 1976). The morphological differences in

Sex Determination and the Development of the Genital Disc

the external terminalia of yakuba–melanogaster and teissieri–melanogaster male hybrids, and those of their parental species, can be either qualitative or quantitative and could be ascribed to four main categories: 1. There were structures in the three species that showed a different shape in their hybrids; for example, the edeagus, the hypandrium, and the lateral lobe. 2. There were structures that in the hybrids showed the morphology of one of the parental species; for example, the dorsal parameters in teissieri– melanogaster resembled those of D. teissieri. 3. There were structures present in the hybrids and in only one of the parental species; e.g., the penis mantle is absent in D. teissieri and present in D. melanogaster and in teissieri–melanogaster hybrids. 4. There were structures absent in the hybrids but present in the two parental species; e.g., the ventral parameres are present in D. melanogaster and D. teissieri but absent in their hybrids. It was speculated that these morphological differences correspond to changes in two basic genetic mechanisms that control the generation of morphological patterns. Qualitative differences, which refer to the presence of absence of a given structure of the pattern, probably reflect a simple genetic basis, in which a single gene is at work (or a combination of a few genes, each contributing a qualitative effect). Quantitative differences, however, are also characterized, because the structures display more or less continuous variation or intermediate phenotypes, suggesting that a polygenic system is at work (each gene in the system has a minor effect and shows additivity) (Sa´ nchez and Santamaria, 1997). The dsx gene is the last gene in the sex determination cascade (Figure 4). The dsxD mutation transforms females into intersexes while having no effect in males (Gowen, 1942; Denell and Jackson, 1972). The XX; dsxD/þ intersexual flies contain both the DsxM (from the dsxD allele) and DsxF (from the dsxþ allele) products (Nagoshi and Baker, 1990). The dsxD allele was introduced into the teissieri– melanogaster hybrids via D. melanogaster males crossed to chimeric D. melanogaster females carrying exclusively D. teissieri germ cells (Sa´ nchez and Santamaria, 1997). The most conspicuous feature of the intersexual terminalia of XX; dsxD/þ D. melanogaster flies is the presence of both male and female genital structures. These are reduced and arranged in two separate genitalia. Some intersexes show a reduction of genital structures to such an

29

extent that only traces of male and/or female genitalia are present. The anal plates are always present and their shape resembles neither the wild-type male nor the normal female. Rather they display an intermediate sexual phenotype (Gowen, 1942; Denell and Jackson, 1972; Epper, 1981). The most salient characteristic of the intersexual terminalia of XX; dsxD/þ teissieri–melanogaster hybrids was the great difference in the inventories of male versus female genital structures: in general terms, the external genitalia of these hybrids could be defined as malelike more than intersexual. There was an almost completely normal set of male genital structures. The analia, however, were intersexual (Sa´ nchez and Santamaria, 1997). As mentioned earlier, the dsx gene must act in concert with another regulatory gene(s) to determine the genital primordia that will develop in each sex. It has been proposed that this additional gene is Abdominal-B (Abd-B) (Sa´ nchez et al., 1997, 2001). To explain the male-like phentoype of the genitalia of XX; dsxD/þ teissieri–melanogaster hybrids, it was speculated that during the evolution of the D. melanogaster and D. teissieri species, genetic changes have occurred in the regulatory genes, such as dsx and/or Abd-B, and/or in the genes controlled by these regulators, which are responsible for the development of the terminalia (Sa´ nchez and Santamaria, 1997). These species-specific variations could be responsible for the morphological changes observed in the terminalia of these species. When the genotypes of the two species are put together within a single hybrid cell, divergent coadapted gene complexes are confronted. This might result in the formation of hybrid patterns different from those of their parental species; i.e., in the production of morphological diversity. 1.1.2.6.2. The genital disc in other dipteran species The expression of pattern-forming genes in the genital disc indicates that this disc is organized like the ventral, leg, and antenna discs (see Chapter 1.9). Evolutionary studies have suggested that the terminalia arose as a modification of a primitive appendage of a common ancestor to all arthropods (Matsuda, 1976). It has also been suggested that Dll expression along the proximal–distal axis arose once in an appendage-bearing ancestor, and has been utilized repeatedly in body wall outgrowths ever since (Panganiban et al., 1997). It can be hypothesized that the Abd-B function has modified this ‘‘ground state’’ in order to allow genitalia differentiation. This modification appears to be concomitant with the tendency towards a reduction of the structures (through the fusion of corresponding segments)

30 Sex Determination and the Development of the Genital Disc

posterior to the 5th abdominal segment. Such reductions have occurred to different extents in males and females, leading to an increase in sexual dimorphism (Crampton, 1942; Matsuda, 1976). Two major features characterize this evolution: a decrease in the number and size of posterior-most tergites and sternites, and the fusion of posterior segments into a single genital disc (Crampton, 1942; Matsuda, 1976). Insects such as Musca (Du¨ bendorfer, 1971) and Calliphora (Emmert 1972a, 1972b), which are more primitive than Drosophila, have two lateral and one median genital discs. The primordium of the lateral discs corresponds to A8, and that of the single median disc to the fusion of A9 to A11 (Figure 14). In females, the lateral discs form the female genitalia, except for the parovaria. The median disc forms the parovaria (A9) and the female analia (A10–A11). In males, the lateral discs produce a reduced eighth tergite. The median disc develops the male genitalia (A9) and the male analia (segments A10–A11) (Figure 14). A further level of fusion has occurred in the Drosophila lineage, where segments A8 to A11 form a single genital disc (Figure 14).

1.1.3. Concluding Remarks and Perspectives Perpetuation by sexual reproduction is the rule within the animal kingdom. The sex determination mechanisms have long been of major interest from a developmental and evolutionary point of view. There is a plethora of mechanisms whereby formation of males and females occurs through a species’ ontogeny. This is more evident in insects, where all types of sex determination mechanisms are represented. The characterization of the genes that control sexual development in Drosophila contributed significantly to the understanding of the molecular strategies behind sex determination, sex behavior, and the development of the germline (see Chapters 1.2 and 1.4). This knowledge paved the way for the study of the sex-determination mechanisms in other insects, so that the understanding of evolution of the sex-determining mechanisms is now possible. This will undoubtedly also contribute to an increased understanding of the evolution of other genetic pathways. In addition, results from studies on the mechanisms that control sexual development, are likely to contribute to the development of novel

Figure 14 Hypothetical evolutionary pathway of the genital disc. Comparison of the genital discs of Musca (upper) and Drosophila (lower). A8–11 stands for abdominal segments 8–11. Strong experimental evidence supports the idea that the anal primordium of the Drosophila genital disc shows a complex organization (see text for explanation). Therefore, it is illustrated that two cell populations, assumed to be derived from segments A10 and A11, respectively, form the anal primordium (the horizontal broken line indicates the different segmental origin). It is further assumed that this organization is already present in Musca. The dotted line running from top to bottom indicates the bilateral symmetry organization of the genital disc.

s0170

Sex Determination and the Development of the Genital Disc

strategies for the control or eradication of harmful insects that fhave a serious detrimental economic impact in agriculture and affect human health. Traditionally, the control of pest insects has been performed by the use of insecticides. Although this strategy is quite effective, it has important disadvantages including the pollution of the environment, a significant risk for human health, and the appearance of insecticide resistance in the pest population. This is the reason why biological control of pest insects is receiving increasing attention. The biological control of insect pests has been carried out by the release of sterilized adults into the pest population, as is the case with the fruit flies. Although this method can be quite effective, there are certain aspects that are undesirable. Only the sterile males contribute positively to pest control, yet both males and females are mass-reared and released into the environment. Although females are sterile, they are undesirable because they can still cause crop damage through oviposition. Consequently, the selective elimination of females would be extremely important for biological control of insect pests. Two potential strategies can accomplish this goal. One of them is based on the selective expression, in females, of a harmful gene, which would be under the control of female-specific regulatory sequences. The second strategy is based on the transformation of females into sterile males or intersexes lacking the ovipositor structure, due to the expression of a gene that causes male instead of female development. Both strategies require the isolation and further characterization of the genes controlling sex determination and/or sex-specific regulatory sequences, as well as the development of vectors (transposable elements) that would allow the formation of transgenic fruit flies (see also Chapters 4.12, 4.13, and 5.12). Drosophila has a single genital disc that produces the terminalia, the most sexually dimorphic structure in the adult. The founder cells are recruited from different segments to form the two genital primordia, female and male, plus the anal primordium that constitute the genital disc. In addition to being a good experimental model for the study of the control of pattern formation, the genital disc is also well suited for the study of morphological evolution. In this respect, two aspects of the genital disc can be highlighted. First, the fact that the female external terminalia are very similar in the Drosophila species. The external male terminalia, however, display great differences. Therefore, the terminalia are a good experimental model for understanding the mechanisms controlling morphogenesis. In

31

addition, they are particularly well suited for studying the causes determining the relative morphological simplicity observed in the female genitalia, and the great morphological variety observed in the male genitalia. The second aspect is related to the origin of the genital disc. The expression of patternforming genes in the genital disc supports the idea that the terminalia arose as the result of the modification of a primitive appendage of a common ancestor to all arthropods. This modification appears to be concomitant with the tendency towards the reduction of structures (through the fusion of corresponding segments) posterior to segment A5. Such reductions have occurred to different extents in males and females, leading to an increase in sexual dimorphism. The knowledge acquired from the study of the Drosophila genital disc allows us to address questions about the organization and development of the genital disc in other insects. This will aid us to understand the evolution of the genital disc and, ultimately, morphological evolution.

Acknowledgments This work was financed by grant BMC2002-02858 awarded to L.S. and grant BMC2002-03839 awarded to I.G. by the Comisio´ n Interministerial de Ciencia y Tecnologı´a (CICYT), and an institutional grant from the Fundacio´ n Areces. N.G. was supported by a research contract I3P from Consejo Supeior de Investigaciones Cientı´ficas (CSIC).

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32 Sex Determination and the Development of the Genital Disc

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1.2 Oogenesis D A Dansereau and P Lasko, McGill University, Montreal, QC, Canada D McKearin, University of Texas, Dallas, TX, USA ß 2005, Elsevier BV. All Rights Reserved.

1.2.1. Introduction 1.2.2. Stem Cells 1.2.2.1. Germline Stem Cells and the Stem Cell Niche 1.2.2.2. Somatic Stem Cells 1.2.2.3. Stem Cell Niche Cell Adhesion 1.2.2.4. Stem Cell Niche Communication 1.2.3. Germline Cyst Formation 1.2.3.1. Ring Canals 1.2.3.2. The Fusome 1.2.3.3. The Fusome Cycle 1.2.3.4. Counting Cystocyte Cell Divisions 1.2.4. Early Oocyte Differentiation and Polarity 1.2.4.1. Establishing the Oocyte and Early Polarity 1.2.4.2. The Spindle Genes, Meiotic Checkpoint, and Patterning 1.2.5. Follicle Cells 1.2.5.1. The Organizing Activity of Polar Cells 1.2.5.2. Polar Cells Transmit Polarity to Adjacent Cysts 1.2.5.3. Determining the Polar Cells 1.2.6. Establishing Embryonic Polarity 1.2.6.1. Oocyte Cytoskeletal Polarity and oskar Localization 1.2.6.2. Dynein and Kinesin: Dueling Motors 1.2.6.3. RNA Recognition Complexes 1.2.6.4. Nurse Cell-Oocyte Cytoplasmic Transfer and Late-Localizing mRNAs 1.2.6.5. Embryonic Anterior/Posterior Polarity 1.2.6.6. Dorsal–Ventral Polarity 1.2.7. Primordial Germ Cells 1.2.7.1. PGC Fate Specification 1.2.7.2. Cell Cycle Control and PGC Migration 1.2.7.3. Transcriptional Repression in PGCs

39 40 40 42 43 43 45 45 46 46 47 49 49 51 53 54 55 55 57 58 58 59 60 62 62 66 66 68 69

1.2.1. Introduction Drosophila have polytrophic ovarioles: each oocyte develops in an egg chamber (follicle), containing a cluster of interconnected germline cells (cyst) surrounded by a layer of mesodermal follicle cells. In principle, whatever applies to Drosophila also broadly applies to many other insects with polytrophic ovarioles (e.g., Anopheles and Bombyx; see also Chapter 1.3). The Drosophila oocyte develops within a cyst of 16 interconnected germline cells, all derived from one cystoblast through mitotic divisions. Egg chambers are assembled in the germarium, a mitotically active zone at the anterior tip of each ovariole. Here, mitotic division of a germline stem cell (GSC) renews the GSC and produces a cystoblast, which undergoes four synchronous mitotic divisions to

generate a 16-cell cyst (Figure 1). Cystocyte cell cycles are rapid and uncoupled from cell growth, leading to a progressive reduction in cell volume. Older cysts migrate posteriorly through the germarium, and are individually surrounded by mesodermal follicle cells, thus forming egg chambers. Follicle cell migration is associated with the flattening and lengthening of the cyst perpendicular to the anterior–posterior (A–P) axis so that it spans the width of the germarium. Each germarium, which can be further subdivided into four consecutive and morphologically and functionally distinct regions (1, 2A, 2B, and 3; Figure 2), typically contains 7–12 developing 16-cell cysts. There are 6–7 egg chambers per ovariole connected to each other by stalks of follicle cells (Figure 2). As

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40 Oogenesis

development proceeds, a rhythmically contracting muscular epithelial sheath pushes egg chambers progressively toward the posterior. Thus, each ovariole can be considered as a linear assembly line for the formation of eggs, with progressively older egg chambers toward the posterior. Seventeen to 20 discrete tubular ovarioles are packaged together in parallel, in each of the two ovaries. Each ovary is surrounded by a network of muscle fibers called the peritoneal sheath and both are connected to a common oviduct with associated accessory sex glands (see Chapter 1.5). As mature eggs pass from the common oviduct into the uterus, they are fertilized by sperm stored in the ventral receptacle and spermathecae. The germline-derived cell types are the GSCs and the cystocytes of individual egg chambers, which develop into nurse cells and the oocyte. Somatic tissue surrounding the egg chambers is derived from the gonadal mesoderm. All other structures of the reproductive system differentiate from the genital imaginal disc (see Chapter 1.1). Oogenesis can been divided into 14 stages based on morphological criteria (King, 1970). Stage 1 refers to a newly formed egg chamber and stage 14 is a mature egg that is ready to be fertilized and oviposited. The mature egg is visibly asymmetric along its A–P and dorsal–ventral (D–V) axes (Figure 2). It has an anterior micropyle and anterior dorsal appendages, a posterior aeropyle, and is more curved on the ventral surface relative to the dorsal one. This polarity is defined and maintained during oogenesis through communication between the follicle cells and the developing nurse cell-oocyte complex.

1.2.2. Stem Cells 1.2.2.1. Germline Stem Cells and the Stem Cell Niche

Stem cells are defined by their ability to self-renew and produce daughters that differentiate. There are

f0005

Figure 1 Cyst divisions in region 1 of the germarium. Oogenesis begins with the asymmetric division of a germline stem cell (GSC) at the anterior tip of germarial region 1. One daughter remains in the stem cell niche and is maintained as a GSC, while the second leaves the stem cell niche and differentiates as a cystoblast (discussed in Section 1.2.2). Each cystoblast divides four times (I–IV) with incomplete cytokinesis to give rise to a cyst of 16 cells connected to each other by cytoplasmic bridges called ring canals (discussed in Section 1.2.3). This leads to the production of a cyst containing only two cells with

four ring canals (marked 4r), one of which becomes the oocyte and undergoes meiosis (discussed in Section 1.2.4.1). Broken arrows denote centriole migration and open arrowheads represent mitotic spindles. During these divisions, a membranous cytoplasmic organelle called the fusome, shown in red, anchors one pole of each mitotic spindle, orienting and synchronizing each cyst division (discussed in Sections 1.2.3.2 and 1.2.3.3). These organelles are spherical in GSCs, where they are called spectrosomes, but grow and fuse to become elongated, branched structures that extend through each ring canal during cyst divisions. The cystocyte that is chosen to become the oocyte might be determined as early as the first cystoblast division, when one daughter inherits more fusome than the other. This initial asymmetry is propagated through subsequent divisions and likely plays a central role in specifying oocyte fate (discussed in Section 1.2.3.2).

s0015

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Oogenesis

41

Figure 2 Drosophila ovarioles and egg chambers. Drosophila ovaries are organized into approximately 17 tubular structures called ovarioles that contain progressively maturing egg chambers. Different developmental stages from a single ovariole are pictured here. Egg chamber development initiates in the germarium at the anterior tip of each ovariole, which can be divided into three sections based on different stages of cyst development. Region 1 is filled with mitotically active cells: germline stem cells (GSCs), somatic stem cells (SSCs), and dividing cystocytes (Sections 1.2.2 and 1.2.3). In region 2, 16-cell cysts change shape to take up the width of the germarium, and the oocyte is determined (discussed in Section 1.2.2.4.1). Cysts are encapsulated by somatic follicle cells in region 2, and a subpopulation of follicle cells between adjacent cysts intercalates in region 3 to form a stalk, causing the newly formed stage 1 egg chamber to bud from the germarium (discussed in Section 1.2.5.1). Oogenesis can be further subdivided into 14 stages based on morphological criteria, where stage 1 refers to a newly formed egg chamber and stage 14 is a mature egg that is ready to be fertilized and oviposited. During stages 1–6, a stable microtubule organizing center (MTOC, marked with a black oval and minus sign in the stage 6 egg chamber pictured here) is present at the posterior of the oocyte (discussed in Section 1.2.4.1.1). It nucleates a single microtubule network (depicted as black lines) emanating through the ring canals into each nurse cell, with most MT minus ends focused in the oocyte. Until the cytoskeleton is reorganized during stages 6–7, mRNAs and proteins are steadily transported toward the MT minus ends, so that they accumulate in the oocyte. Major cytoskeletal reorganization occurs during stages 7–8, after which the posterior MTOC is no longer detected. A reorganized cytoskeleton emerges with the bulk of the MT network at the anterior, creating a higher density of MT in the anterior of the oocyte than the posterior. MT minus ends appear to be nucleated along the anterior and lateral cortices, with MT plus ends pointing into the center of the oocyte (discussed in Section 1.2.6.1 and portrayed in the stage 10 egg chamber drawn here). This cytoskeletal organization is required to direct the posterior localization of osk mRNA, the anterior localization of bcd mRNA, and the dorsal-anterior relocalization of the oocyte nucleus and grk mRNA. The mature egg (bottom drawing) is visibly asymmetric along its anterior–posterior (A–P) and dorsal–ventral (D–V) axes. It has an anterior micropyle and anterior dorsal appendages, a posterior aeropyle, and the egg is more curved on the ventral than on the dorsal surface.

two types of stem cells in the ovary: GSCs and somatic follicle stem cells (SSCs, Figure 3; Margolis and Spradling, 1995; Spradling et al., 1997, 2001). The anterior of each developing germarium hosts on average 2 or 3 GSCs, each of which is located adjacent to two or three somatic cap cells (Forbes et al.,

1996a; Spradling et al., 1997; Xie and Spradling, 2000). Anterior to the cap cells in each ovariole are about six terminal filament (TF) cells which are arranged in a single-file array resembling a stack of pancakes. To the posterior of the cap cells and surrounding the GSCs and newly formed cystocytes are

42 Oogenesis

f9000

Figure 3 The ovarian germline and somatic stem cell niche. The germline stem cells (GSCs, yellow) and somatic stem cells (SSCs, red) are maintained in a somatic niche in germarial region 1, illustrated here. The stem cell niche is composed of terminal filament cells (TFs, blue), cap cells (blue), and inner germarial sheath cells (IGS, pink). TF and cap cells express Yb, Piwi, and the signaling molecules Dpp, Wg, and Hh, which act together to coordinate the two stem cell populations. GSCs maintain close contact with the cap cells through adherens junctions. Dpp signaling activity is restricted to GSCs where it maintains GSC fate by repressing Bam transcription, thereby preventing differentiation. When a GSC divides, one daughter maintains contact with the cap cells and remains a GSC, while the other daughter (the cystoblast – CB) escapes the niche and activates Bam expression (BamON, shown in green). Bam is required to execute the cystoblast division program of four incomplete divisions, forming 16-cell cysts that no longer express Bam (BamOFF). In addition to intercellular signaling pathways, cell-autonomous factors like the translational repressors Nanos and Pumilio (Nos/Pum) are required in the germline for the maintenance of GSCs. SSCs are located at the boundary between germarial regions 1 and 2, and serve as the source of prefollicle cells (brown) that will encapsulate individual cysts. Adherens junctions between the SSCs and IGS cells retain SSCs in their niche. Hh and Wg are the only known signals for maintaining SSCs. Yb is expressed in the TF and cap cells where it activates the Piwi pathway for GSC maintenance, and the Hh pathway for both GSC and SSC maintenance. Molecular targets of Piwi, Hh, and Wg in stem cell maintenance have not been identified.

the inner germarial sheath (IGS) cells (Figure 3; Godt and Laski, 1995; Chen et al., 2001). The TF cells, cap cells, and IGS cells constitute the stem cell niche – the cellular microenvironment that regulates, induces and supports the development of stem cells (Figure 3; review: Spradling et al., 2001). The TF and somatic cap cells are the likely sources of several important signaling molecules, and the survival of GSCs is dependent on their close association with these cells. Niche function also depends on the continued presence of stem cells, since empty niches eventually degenerate (Xie and Spradling, 2000). Thus, the GSCs and the stem cell niche function together to maintain the size of the stem cell pool, and allow the regular production of new egg chambers (Spradling et al., 2001; Lin, 2002; Gonzalez-Reyes, 2003; Kai and Spradling, 2003). Cap cells and TF cells can be visualized with a hedgehog-lacZ enhancer trap and the IGS cells can be identified by patched-lacZ expression (Forbes et al., 1996a). The GSCs can be recognized because of their large size, location, and the presence of a cytoplasmic organelle called a spectrosome that

is invariably anchored to the cap cell-GSC contact site (Figure 1; Lin et al., 1994; de Cuevas and Spradling, 1998). 1.2.2.2. Somatic Stem Cells

Continued generation of follicle cells is ensured by the maintenance of two or three somatic stem cells that are found at the surface of the germarium at the 2A/2B boundary (Figures 2 and 3). The immediate progeny of the SSCs, the follicle cell progenitors, divide to generate different classes of follicle cells that encapsulate each germline cyst. No positive marker for the SSCs has been described so far and they cannot be recognized by morphology or position, thus complicating their study. The SSCs have been identified as the only mitotically active somatic cells that remain in the germarium and continue to divide after being genetically marked (Margolis and Spradling, 1995). Zhang and Kalderon (2001) noted that the anterior-most mitotically active somatic cells fail to stain with Fasciclin III (FasIII) antibodies, which mark differentiating follicle cells, and have suggested that these are the SSCs.

s0020

p0040

Oogenesis

1.2.2.3. Stem Cell Niche Cell Adhesion

During late larval and early pupal development, the somatic mesoderm starts to differentiate, dividing the ovary into individual ovarioles. The TFs form during the second half of the third larval instar. Adherens junctions form between the cap cells and the newly established GSCs during early pupation (Figure 3). After the formation of the TFs and the cap cells, the anterior primordial germ cells (PGCs) are selected to become GSCs while the rest of the PGCs are thought to enter directly the cyst differentiation pathway (Bhat and Schedl, 1997; Song et al., 2002; Zhu and Xie, 2003). Cell adhesion mediated by DE-Cadherin plays an important role during late larval development in the recruitment of PGCs to the stem cell niche where they become GSCs (Song and Xie, 2002). Without DE-Cadherin, germline cells are not efficiently incorporated into the niche as stem cells. Contact between germline cells and newly differentiated cap cells seems to initiate the establishment of stable adherens junctions between them. The adherens junction components, DE-Cadherin and Armadillo (Arm; b-Catenin), are concentrated at the interfaces between the cap cells and GSCs. These adherens junctions are essential to hold the GSC in the niche and, therefore, maintain its stem cell identity (Figure 3; Lin and Spradling, 1993; Xie and Spradling, 1998). Removal of DE-Cadherin or Arm from the GSCs allows them to leave the stem cell niche and differentiate (Song and Xie, 2002). The GSCs divide to produce cystoblasts that differentiate into germline cysts (Figure 1). GSCs divide along the A–P axis of the germarium so that the anterior daughter remains anchored to the cap cells and retains stem cell identity. The other daughter, now one cell diameter from the cap cells, differentiates as a cystoblast and begins oogenesis (Lin and Spradling, 1997). Stem cells and cystoblasts are connected by a transient ring canal (Carpenter, 1981; de Cuevas and Spradling, 1998). Before either cell divides again, the ring canal shrinks and the connection between the two cells is severed. The GSCs are randomly lost to differentiation, with a half-life of about 4–5 weeks (Margolis and Spradling, 1995; Xie and Spradling, 1998). When a GSC is lost, the remaining stem cells can divide perpendicular to the A–P axis so that both daughters remain attached to the cap cells (Xie and Spradling, 2000; Zhu and Xie, 2003). This division pattern suggests that symmetrical GSC division replenishes any vacant niche spaces. As with GSCs, Cadherin-mediated cell adhesion is probably required for anchoring SSCs to their stem cell niche (Song and Xie, 2002, 2003). The SSCs

43

interact directly with the IGS cells, which may function as the SSC niche. DE-Cadherin and Arm accumulate at the border between SSCs and IGS cells, and removal of DE-Cadherin or Arm from the SSCs allows them to move away from their niche and differentiate. While the asymmetry of GSCs divisions is clearly marked by asymmetric cytoplasmic inheritance of the spectrosome (Section 1.2.3.3), the division of a SSC is not necessarily asymmetric: the different behavior of the SSC daughters can be explained if the stem cell niche has a limited capacity and one daughter cell must leave the niche after division. 1.2.2.4. Stem Cell Niche Communication

Signals emanating from the GSC niche are required for the maintenance of both the GSCs and the SSCs. TF and cap cells express the genes Yb, piwi, decapentaplegic (dpp), wingless (wg), and hedgehog (hh), which are part of an intercellular signaling network that coordinates the two stem cell populations (Figure 3). Intrinsic factors, including the Decapentaplegic (Dpp) receptor, Pumilio (Pum) and Nanos (Nos), are required in the GSCs for maintenance.1 1.2.2.4.1. Dpp Dpp, the Drosphila homolog of TGF-b/BMP, is secreted by the somatic cells in the stem cell niche and received by the GSCs, where the Dpp signaling pathway is activated (Xie and Spradling, 1998, 2000; Song et al., 2004). The dpp mRNA is expressed in the cap cells, and the Dpp receptor, and downstream proteins in the signaling pathway, are required in the GSCs. Overexpressing dpp in somatic cells is sufficient to induce more GSCs, and misexpression of dpp or ectopic activation of the Dpp pathway in GSCs results in germ cell overproliferation, indicating that Dpp signaling stimulates GSC division and represses differentiation of the daughter cells. GSCs lacking Dpp receptors are replaced by wild-type germline cells, which come to lie next to the cap cells and behave as GSCs, indicating that the Dpp signal maintains GSCs in a niche that is able to program newly introduced cells to become stem cells. In fact, Dpp appears to be sufficient to cause four- and eight-cell germline cystocystes to de-differentiate and repopulate the GSC niche (Kai and Spradling, 2004). The activity of dpp maintains GSCs in their niche because it represses the expression of bag of marbles (bam) (Chen and McKearin, 2003b; Song et al., 2004), a factor required to resume a program of 1

An appendix at the end of this chapter includes a list of gene names, their abbreviations, and a short description of their functions.

p0065

p0070

44 Oogenesis

cyst cell division and differentiation (McKearin and Spradling, 1990; McKearin and Ohlstein, 1995). The bam gene is expressed in the cystoblasts but not in the GSCs (Figure 3; McKearin and Ohlstein, 1995; Chen and McKearin, 2003b). Overexpression of bam in GSCs effectively eliminates the GSC fate, similar to the loss of dpp signal transduction in the GSCs (Ohlstein and McKearin, 1997). Significantly, loss of dpp expression in the stem cell niche quickly allows the expression of bam in the GSCs (Song et al., 2004). Conversely, broad expression of dpp, directly prevents bam transcription and blocks the differentiation of GSC daughter cells (Chen and McKearin, 2003a). bam appears to be the principal Dpp target because ectopic Bam expression is sufficient to restore differentiation to Dpp-overexpressing cells, and because the bam promoter contains a transcriptional silencer element that binds the Mothers against Dpp (Mad) and Medea proteins (Chen and McKearin, 2003a, 2003b; Song et al., 2004). The simplest explanation for these results is that Dpp functions as a short-range signal that promotes GSC survival by repressing bam expression specifically in the GSCs and preventing their differentiation as cystoblasts. 1.2.2.4.2. Nanos and Pumilio The RNA binding proteins Nanos (Nos) and Pumilio (Pum) are required in the germline for the maintenance of GSCs (Lin and Spradling, 1997; Forbes and Lehmann, 1998; Bhat, 1999; Parisi and Lin, 1999). Ovaries mutant for nos or pum have very few germ cells, and the number of germ cells decreases with age, suggesting that these two genes are required for GSC maintenance (Forbes and Lehmann, 1998; Bhat, 1999; Wang and Lin, 2004). nos is expressed at a high level in all PGCs in the third instar larval ovary. If nos function is removed during this period, a large proportion of PGCs, including those in contact with the stem cell niche, prematurely differentiate into cysts (Wang and Lin, 2004). These results indicate that nos activity maintains GSC self-renewal by preventing differentiation. The ovarian phenotype of a nos pum double mutant is similar to either nos or pum alone (Wang and Lin, 2004), suggesting they act as partners in GSC maintenance. As a Nos/Pum complex acts as a translational repressor in later stages of oogenesis (review: Parisi and Lin, 2000), these molecules are presumed to maintain GSC fate by repressing translation of one or more mRNAs required for differentiation. 1.2.2.4.3. Hedgehog and Wingless Hedgehog (Hh) and Wingless (Wg), expressed by the TF and cap cells, are the only known signals required for the

maintenance of somatic stem cells (SSCs) (Forbes et al., 1996a, 1996b; Zhang and Kalderon, 2000, 2001; King et al., 2001; Song and Xie, 2003). Loss of Wg signal transduction or constitutive activation of the Wg pathway in SSCs results in their rapid loss, suggesting that correct intermediate levels of Wg signaling are important for proper regulation of SSCs. In addition, constitutive Wg signaling causes overproliferation and increased differentiation of polar cells and stalk cells (Section 1.2.5.3), causing defects in egg chamber budding and abnormal egg chamber morphology. Hh also regulates directly SSC maintenance and proliferation. Unlike wg, however, overexpression of hh induces SSC proliferation; like wg, a reduction in Hh signaling causes rapid SSC loss and arrests egg chamber budding. Hh may also have a partially redundant function in GSC maintenance, since loss of hh causes loss of about 20% of the GSCs, while its overexpression stimulates a slight increase in the number of GSCs. The hypothesis that Hh positively regulates GSC survival is further supported by the observation that hh overexpression restores GSC maintenance to both piwi and Yb mutants (see below). 1.2.2.4.4. Piwi and Yb Two other genes known to be involved in maintaining stem cell survival are piwi (Cox et al., 1998, 2000), and Yb (King and Lin, 1999; King et al., 2001). Overexpression of either gene allows the specification of extra GSCs, while loss of either leads to the differentiation of GSCs into a few dividing cystoblasts, thus depleting the germline. Both are expressed and autonomously required in the TF cells and cap cells. piwi is also expressed in the germline, but is required specifically in the somatic cells for GSC maintenance. In Yb mutant germaria, piwi expression is specifically eliminated in the TF and cap cells, suggesting that Yb acts through piwi to maintain GSCs. Piwi is likely to be of fundamental importance as it is the founding member of a conserved class of proteins (the Piwi, Argonaute, or PPD family) that function in stem cell maintenance in both the animal and plant kingdoms (reviews: Benfey, 1999; Cerutti et al., 2000). The step of gene expression on which Piwi may exert regulation is unclear, as Piwi-related proteins have been implicated in several DNA- and RNA-related processes including RNA interference (Smulders-Srinivasan and Lin, 2003). Yb is a positive regulator of both hh and piwi expression. Since hh is required primarily for SSC maintenance, and piwi is required primarily for GSC maintenance, Yb appears to coordinately regulate SSCs, through hh, and GSCs, through piwi, hh, and an uncharacterized signal (King et al., 2001). Yb

Oogenesis

regulates SSC proliferation by autonomously activating hh expression in the stem cell niche. Ectopic hh expression can induce SSC proliferation and partially restore GSC maintenance to Yb mutants, suggesting that Yb acts upstream of hh to regulate both stem cell populations. While piwi and hh are both involved in maintaining GSCs, it is believed that they function in separate parallel pathways, with piwi having the more important role in normal development. However, hh overexpression increases the number of GSCs even in piwi mutants, therefore ectopic hh is able to rescue the GSC loss phenotype of piwi. It is unlikely that this effect comes about through an Hh-dependent increase in the capacity of the niche, as overexpression of dpp does not rescue the effects of piwi on GSC number (King et al., 2001).

1.2.3. Germline Cyst Formation The mitotically active region at the anterior of each ovariole is termed the germarium. It can be divided into four regions organized along the A–P axis (1, 2A, 2B, and 3) that correspond to maturing cyst stages (Figures 2 and 3). Region 1 contains 2–3 GSCs, cystoblasts and interconnected cysts of 2, 4, or 8 cells. Two or three GSCs occupy the apical position in each ovariole and are in close contact with several anterior somatic cells (Figure 3). GSC division produces a new GSC that remains in the position of the mother cell, and a second posterior daughter that develops as a cystoblast (King, 1970). The cystoblast divides four times with incomplete cytokinesis at each division, producing a syncytial cluster of 16 interconnected cystocytes (the cyst; Figure 1). Comparative analyses of germ cell development suggest that similar cell lineages arise during gametogenesis in diverse organisms (de Cuevas and Spradling, 1998; Pepling et al., 1999). Cytokinesis is incomplete, since cleavage furrows are arrested and become stabilized as ring canals (see below). Similar ring canals form in most germ cells, including those from vertebrate models, and may serve to synchronize germ cell maturation (Pepling et al., 1999). Prefollicle cells, produced by 2–3 SSCs that reside on the side of the germarium at the regions 2A/2B (Figure 3), surround the germline cysts to form an epithelium and separate cysts from one another (Figures 2 and 3). The newly formed follicles or egg chambers change from spherical to elliptical so that they span the entire width of the germarium. The oocyte is centrally located in the cyst at this time. In regions 2B and 3, the newly specified oocyte takes up a posterior position as the cyst again

45

changes shape to become spherical as it buds from the germarium as a stage 1 egg chamber (Figure 2). Under controlled conditions, it takes 3 days for a cystoblast to develop into a 16-cell cyst (King, 1970), and 7 days to reach stage 1 egg chamber (Wieschaus and Szabad, 1979). Then development of an egg progresses rapidly and is complete within 3 additional days (Lin and Spradling, 1993). 1.2.3.1. Ring Canals

During oogenesis of many insects, including Drosophila, cytoplasmic bridges called ring canals allow the flow of materials among the nurse cells and the oocyte. Ring canals can be visualized by staining with fluorescently labeled phalloidin, which binds to filamentous Actin, or with antibodies directed against ring canal components. The division pattern of the germline cyst leads to only two cells with four ring canals (Figure 1). The ring canal connecting the two daughter cells of the first division is the largest in diameter, and those arising at the second, third, and fourth divisions are each smaller than the previous ones. Ring canals have an electronopaque outer ring associated with membranes and an electron-dense inner rim of Actin, the Hu-Li Tai Shao ring canal (Hts-RC) protein, and Kelch. Their assembly involves the sequential addition of proteins that transform the arrested cleavage furrow into a stable intercellular bridge (de Cuevas and Spradling, 1998). Although the inner rim is established coordinately with the degeneration of the fusome (see below) and the addition of Actin to the ring canal, the fusome is not required for arrest of the contractile ring or to initiate ring canal formation (Section 1.2.3.2; Lin et al., 1994). An outer-rim component that is recognized by anti-phosphotyrosine antibodies begins to mark ring canals in the early germarium, as the cleavage furrows arrest in the final round of cyst mitosis (Robinson et al., 1994). Next, the electron-dense inner ring is formed as Cheerio (Filamin), then Actin and Hts-RC, are added starting in region 2A (Yue and Spradling, 1992; Theurkauf et al., 1993; Robinson et al., 1994). Female sterile cheerio mutants fail to recruit Hts-RC and Actin to the ring canals (Yue and Spradling, 1992; Robinson et al., 1997). Kelch is the last known component to be added to the ring canals and it is added only after the Actin filament structure of the ring canals is complete (Tilney et al., 1996). After Kelch is added, it is required through oogenesis for the continued expansion of the ring canals. kelch mutants initially form normal ring canals, which then become disorganized, resulting in female sterility due to transport blockage from the nurse cells into

46 Oogenesis

the oocyte (Section 1.2.6.4; Xue and Cooley, 1993; Robinson and Cooley, 1997). 1.2.3.2. The Fusome

Reconstruction of the GSC, cystoblast, and cystocyte divisions by electron microscopy (EM) and confocal microscopy demonstrated that mitotic spindles are coordinately oriented during these divisions (King, 1970; Lin and Spradling, 1995). This is accomplished by anchoring one spindle pole of each dividing cell to a complex organelle termed the fusome (Figure 1; McGrail and Hays, 1997). Fusomes can be found in many insect germ cells (Bu¨ ning, 1994), and similar organelles may exist in mammalian lymphocytes (Gregorio et al., 1992; McKearin, 1997), and germ cells (Pepling and Spradling, 1998). The organelles are spherical in GSCs and cystoblasts but grow to become elongated, branched structures that extend through each ring canal as cysts form (Figure 1). Mutations in genes encoding fusome proteins, such as hts or a-spectrin, block fusome formation and cause premature termination of cyst divisions (Lin et al., 1994; de Cuevas et al., 1996). Thus, fusome integrity is essential for proper execution of cyst divisions. In the absence of fusomes, cystocyte spindles are disoriented, mitoses are not synchronized, and cystocytes usually do not complete four full divisions, emerging with unusual number of cystocytes such as 3, 5, or 6. Cysts are approximately of normal size because each cystocyte assumes a volume greater than the average wild-type cystocyte volume. Although cysts containing fewer than 15 nurse cells can complete oogenesis (Mata et al., 2000; D. McKearin, unpublished data), females carrying fusome-inactivating mutations are almost

always sterile because cysts fail to form an oocyte. This observation reveals a second role for fusomes in cyst formation – oocyte determination depends on the fusome, probably because it establishes polarity within the cyst (Section 1.2.4.1). Genetic and immunolocalization studies in Drosophila have identified a large group of proteins found associated with fusomes (Table 1). Our current view of the organelle is that membrane skeleton proteins form a meshwork within which membranous cisternae are embedded. Ultrastructural features suggest that fusome cisternae represent a germ cell-specific modification of the endoplasmic reticulum (ER) (Bu¨ ning, 1994). The finding that an ER-associated protein, Ter94, is a fusome component was consistent with this hypothesis. Furthermore, green fluorescence protein (GFP) that carries ER-targeting signals localizes to the fusome (N. Liu and D. McKearin, unpublished), adding additional support to the idea that the fusome is the site of ER assembly in germ cells. 1.2.3.3. The Fusome Cycle

As indicated above, an asymmetric GSC division produces another GSC and a cystoblast, while asymmetric cystocyte divisions produce 15 nurse cells and a single oocyte. The intrinsic factors that underlie the asymmetric fates of GSC and cystocyte divisions are largely unknown but some features of the asymmetry are clear. Unequal fusome distribution is one prominent manifestation of intrinsic asymmetry, since the future cystoblast inherits a smaller fraction of fusome material than the GSC (Figure 1) (Deng and Lin, 1997; de Cuevas and Spradling, 1998). During cyst divisions, the oldest cystocyte always retains the largest fraction of

Table 1 Fusome proteins Protein

Function

Mutant phenotype

Reference

a-Spectrin b-Spectrin Ankyrin Actin Bag of marbles Cyclin A Dynein Encore Hu-Li Tai Shao KLP61 Lis-1 Orbit

Membrane skeleton Membrane skeleton Scaffold or linker protein Cytoskeleton structure, diverse Novel CDK component Motor Cullin-binding protein Adducin related, cytoskeleton Kinesin-like motor Lissencephaly ortholog Microtubule binding protein, CLASP ortholog Microtubule binding protein Cytoskeletal linker protein ER assembly, vesicle fusion

No fusome No fusome n.d. n.d. Reticulum disrupted Cell lethal Disoriented mitoses >16 cystocytes No fusome Disoriented mitoses 3.0  10
5 5.0  10
7 7.5  10
6 1.4  10
5 1.2  10
6 >10
4 3.5  10
6 5.3  10
6 1.3  10
7 1.6  10
7 7.0  10
7 1.6  10
6 2.5  10
7 4.4  10
6 1.7  10
4 2.2  10
7

9a,14a-Epoxy-20-hydroxyecdysone 9b,14b-Epoxy-20-hydroxyecdysone 9a,14a-Epoxy-20-hydroxyecdysone 2,3-acetonide 25-Fluoropodecdysone B 25-Fluoropolypodine B 25-Fluoroponasterone A

1.4  10
6 2.2  10
7 >10
4 7.2  10
9 4.7  10
8 5.1  10
9

25-Deoxypolypodine B 2-Deoxypolypodine B 3-glucoside 24,25-Didehydrodacryhaninansterone 25,26-Didehydrodacryhainansterone 25,26-Didehydroponasterone A 22,25-Dideoxyecdysone 2,22-Dideoxy-20-hydroxyecdysone 22,23-Di-epi -geradiasterone Dihydropoststerone (two isomers) Dihydropoststerone 2,3-acetonide (20S )Dihydropoststerone 2,3,20-tribenzoate (20R )Dihydropoststerone 2,3,20-tribenzoate (5a-H)Dihydrorubrosterone (5b-H)Dihydrorubrosterone Dihydrorubrosterone 17b-acetate 9,20-Dihydroxyecdysone 20,26-Dihydroxyecdysone (podecdysone C) (22S )20-(2,20 -dimethylfuranyl)Ecdysone (22R )20-(2,20 -dimethylfuranyl)Ecdysone Ecdysone

Ecdysone 2,3-acetonide Ecdysone 6-carboxymethyloxime Ecdysone 22-hemisuccinate Ecdysone 22-myristate 24-epi -Abutasterone 24-epi -Castasterone 22-epi -Ecdysone 3-epi -20-Hydroxyecdysone (coronatasterone)

Reference

Ravi et al. (2001) Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Harmatha et al. (2002) Dinan et al. (1999a), Ravi et al. (2001), Harmatha et al. (2002) Dinan et al. (1999a), Ravi et al. (2001) Odnikov et al. (2002) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Cle´ment et al. (1993), Dinan et al. (1999a, 2003), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Ravi et al. (2001) Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Cle´ment et al. (1993), Harmatha and Dinan (1997) Dinan et al. (1999a, 2003), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (1999a) Cle´ment et al. (1993) Cle´ment et al. (1993) Dinan et al. (unpublished data) Ravi et al. (2001), Hormann et al. (2003) Dinan et al. (1999a) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a, 2003), Ravi et al. (2001) Ravi et al. (2001) Odnikov et al. (2002) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Harmatha et al. (2002) Harmatha and Dinan (1997), Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (1999a, 2003), Ravi et al. (2001) Continued

208 Ecdysteroid Agonists and Antagonists

Table 1 Continued Ecdysteroid a

Geradiasterone 2b,3b,9a,20R,22R,25-Hexahydroxy-5b-cholest-7,14dien-6-one 28-Homobrassinolide 14a-Hydroperoxy-20-hydroxyecdysone 5b-Hydroxyabutasterone 25-Hydroxyatrotosterone A 25-Hydroxyatrotosterone B 24-Hydroxycyasterone 25-Hydroxydacryhainansterone 5b-Hydroxy-25,26-didehydroponasterone A 20-Hydroxyecdysone

EC50 (M )
7

4.0  10 1.8  10
7

Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data)

>10
4 1.6  10
8 2.3  10
8 1.0  10
5 5.0  10
5 2.0  10
6 2.6  10
7 6.3  10
8 7.6  10
9 7.5  10
9

Dinan et al. (1999a) Harmatha et al. (2002) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003) Harmatha and Dinan, 1997 Harmatha and Dinan, 1997 Dinan et al. (unpublished data) Bourne et al. (2002) Dinan et al. (1999a), Ravi et al. (2001) Cle´ment et al. (1993) Harmatha and Dinan 1997; Dinan et al. (1999a), 2003; Ravi et al. (2001); Harmatha et al. (2002), Bourne et al. (2002) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Ravi et al. (2001)

20-Hydroxyecdysone 2,3-acetonide 20-Hydroxyecdysone 20,22-acetonide 20-Hydroxyecdysone 22-benzoate 20-Hydroxyecdysone 6-carboxymethyloxime 20-Hydroxyecdysone dimer 20-Hydroxyecdysone 2-b-D-glucopyranoside

3.3  10
6 4.0  10
7 4.7  10
7 4.0  10
6, 2.0  10
7 1.5  10
7 3.0  10
7 4.3  10
9 Inactive 2.1  10
7 2.0  10
5

20-Hydroxyecdysone 3-b-D-glucopyranoside

1.3  10
5

20-Hydroxyecdysone 22-b-D-glucopyranoside

4.7  10
5

20-Hydroxyecdysone 25-b-D-glucopyranoside

8.5  10
6

20-Hydroxyecdysone 2-hemisuccinate 20-Hydroxyecdysone 22-hemisuccinate 20-Hydroxyecdysone 2,3-monoacetonide 20-Hydroxyecdysone 20,22-monoacetonide 20-Hydroxyecdysone 6-oxime 20-Hydroxyecdysone 2,3,22-triacetate 20-Hydroxyecdysone 3b-D-xylopyranoside (limnantheoside A) 6b-Hydroxy-20-hydroxyecdysone 6a-Hydroxy-20-hydroxyecdysone 26-Hydroxypolypodine B 5b-Hydroxystachysterone C (25R/S )-Inokosterone (25R )-Inokosterone (25S )-Inokosterone Inokosterone 26-hemisuccinate Integristerone A

8.3  10
7 Inactive 1.5  10
7 3.0  10
7 2.2  10
6 >1.6  10
5 1.6  10
6

(5a-H)20-Hydroxyecdysone 20-Hydroxyecdysone 2-acetate 20-Hydroxyecdysone 3-acetate 20-Hydroxyecdysone 22-acetate

20-iso -Ecdysone 20-iso -22-epi-Ecdysone iso-Homobrassinolide Isostachysterone C (D25(26)) Kaladasterone Ketodiol 5a-Ketodiol Leuzeasterone Limnantheoside C

Reference

1.7  10
7 2.0  10
6 4.8  10
7 3.5  10
8 1.1  10
7 1.5  10
7 2.7  10
7 3.8  10
6 1.8  10
7, 8.3  10
7, 2.0  10
7 1.0  10
4 1.0  10
4 >10
5 9.2  10
9, 4.2  10
9 3.4  10
7 Inactive >2.3  10
5 1.7  10
8 1.3  10
6

Dinan et al. (unpublished data) Dinan et al. (unpublished data) Ravi et al. (2001) Cle´ment et al. (1993) Harmatha et al. (2002) Harmatha and Dinan, 1997, Dinan et al. (1999a), Ravi et al. (2001) Harmatha and Dinan, 1997, Dinan et al. (1999a), Ravi et al. (2001) Harmatha and Dinan, 1997, Dinan et al. (1999a), Ravi et al. (2001) Harmatha and Dinan, 1997, Dinan et al. (1999a), Ravi et al. (2001) Cle´ment et al. (1993) Cle´ment et al. (1993) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (1999a) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Cle´ment et al. (1993) Dinan et al. (1999a), Ravi et al. (2001) Cle´ment et al. (1993), Dinan et al. (2003) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Cle´ment et al. (1993) Dinan et al. (unpublished data)

Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a) Dinan et al. (unpublished data) Bourne et al. (2002) Dinan et al. (1999a) Dinan et al. (1999a) Dinan et al. (unpublished data) Meng et al. (2001b)

Ecdysteroid Agonists and Antagonists

209

Table 1 Continued Ecdysteroid a

EC50 (M )
8

Makisterone A

1.3  10

Makisterone C

2.0  10
7

Malacosterone

9.0  10
6

24-Methylecdysone (20-deoxymakisterone A) 14-Methyl-12-en-15,20-dihydroxyecdysone 14-Methyl-12-en-shidasterone Muristerone A

2.1  10
6 2.3  10
6 4.0  10
6 2.2  10
8

29-Norcyasterone 29-Norsengosterone Paxillosterone Paxillosterone 20,22-para-hydroxybenzylidene acetal 14-Perhydroxy-20-hydroxyecdysone Pinnatasterone Podecdysone B Polypodine B

1.2  10
8 1.3  10
7 4.2  10
7 3.0  10
7 1.6  10
8 4.0  10
7 1.2  10
5 1.0  10
9

Polyporusterone B Ponasterone A

2.1  10
9 3.1  10
10

Ponasterone A 6-carboxymethyloxime Ponasterone A dimer Ponasterone A 2-hemisuccinate Ponasterone A 22-hemisuccinate Ponasterone A 3b-D-xylopyranoside (limnantheoside B) Poststerone Poststerone 20-dansylhydrazine Pterosterone (rhapontisterone) Punisterone Rapisterone B Rapisterone C Rapisterone D Rubrosterone Sengosterone Shidasterone (stachysterone D) Sidisterone Sileneoside A (20-hydroxyecdysone 22-galactoside) Sileneoside C (integristerone A 22-glactoside) Sileneoside D (20-hydroxyecdysone 3-galactoside) Sileneoside E (2-deoxyecdysone 3-galactoside; blechnoside A) Stachysterone B Stachysterone C 2,14,22,25-Tetradeoxy-5a-ecdysone 2a,3a,22S,25-Tetrahydroxy-5a-cholestan-6-one 2b,3b,20R,22R-Tetrahydroxy-25-fluoro-5b-cholest8,14-dien-6-one 2b,3b,6a-Trihydroxy-5b-cholestane 2b,3b,6b-Trihydroxy-5b-cholestane Turkesterone

2.0  10
10 7.5  10
6 4.6  10
8 3.1  10
7 5.0  10
7 1.5  10
5

Reference

Cle´ment et al. (1993), Harmatha and Dinan (1997), Dinan et al. (1999a, 2003), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003) Harmatha and Dinan (1997), Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Cle´ment et al. (1993), Dinan et al. (1999a, 2003), Ravi et al. (2001); Harmatha et al. (2002), Bourne et al. (2002) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001) Harmatha and Dinan (1997) Harmatha and Dinan (1997) Harmatha et al. (2002) Dinan et al. (unpublished data) Harmatha et al. (2002), Bourne et al. (2002) Harmatha and Dinan (1997), Dinan et al. (1999a, 2003), Ravi et al. (2001) Dinan et al. (unpublished data) Cle´ment et al. (1993), Harmatha and Dinan (1997), Dinan et al. (1999a, 2003), Ravi et al. (2001) Dinan et al. (unpublished data) Cle´ment et al. (1993) Dinan et al. (unpublished data) Cle´ment et al. (1993) Cle´ment et al. (1993) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003)

ca. 2  10
5 2.0  10
5 6.0  10
6 2.0  10
9 8.3  10
7 2.3  10
7 3.9  10
7 1.0  10
9 >10
4 9.0  10
8 1.5  10
6, 4.0  10
6 4.3  10
6 4.1  10
5 1.0  10
4 3.0  10
5 Inactive

Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (unpublished data)

8.2  10
8 1.4  10
8 7.5  10
9 >2.5  10
5 >10
4 7.2  10
9

Bourne et al. (2002) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data) Dinan et al. (1999a) Dinan et al. (1999a) Dinan et al. (1999a), Ravi et al. (2001)

>2.4  10
5 >2.4  10
5 1.3  10
6

Dinan et al. (1999a) Dinan et al. (1999a) Cle´ment et al. (1993), Dinan et al. (1999a, 2003), Ravi et al. (2001), Bourne et al. (2002) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (unpublished data)

3.0  10
7 8.0  10
7

Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (2003) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003) Dinan et al. (unpublished data) Dinan et al. (unpublished data) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001), Hormann et al. (2003) Dinan et al. (1999a), Ravi et al. (2001) Dinan et al. (1999a), Ravi et al. (2001)

Continued

210 Ecdysteroid Agonists and Antagonists

Table 1 Continued Ecdysteroid a

Turkesterone 2-acetate Turkesterone 11a-acetate Turkesterone 22-acetate Turkesterone 11a-arachidate Turkesterone 11a-butanoate Turkesterone 11a-decanoate Turkesterone 2,11a-diacetate Turkesterone 2,22-diacetate Turkesterone 11a-hexanoate Turkesterone 11a-laurate Turkesterone 11a-myristate Turkesterone 11a-propionate Viticosterone E a

EC50 (M )
6

5.2  10 4.0  10
6 6.0  10
7 2.2  10
5 4.0  10
5 1.1  10
6 2.0  10
4 1.0  10
3 2.2  10
5 3.4  10
6 1.3  10
6 8.3  10
6 1.0  10
7

Reference

Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (unpublished data) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (2003) Dinan et al. (1999a), Ravi et al. (2001)

For structures of individual ecdysteroids, refer to http://www.ecdybase.org.

Figure 5 The structures of brassinosteroids showing ecdysteroid (ant)agonist activity.

in the same species (Richter and Adam, 1991). Spindler et al. (1992) examined the effects of 22S,23S-homobrassinolide and 22S,23S-homocastasterone on an ecdysteroid-responsive Chironomus

tentans cell line and found that both were weak agonists in terms of their ability to induce ecdysteroid-specific responses and that both competed weakly for specifically-bound [3H]PoA in cell-free

Ecdysteroid Agonists and Antagonists

extracts of the C. tentans cells. Assessment of 10 brassinosteroids in the D. melanogaster BII bioassay revealed no agonist or antagonist activity even at their limit of solubility (10
4 or 10
3 M) (Dinan et al., 2001d). In Spodoptera littoralis, 24-epibrassinolide and 24-epicastasterone can compete specifically-bound [3H]PoA with Kd values of 3.7 and 2.0 mM, respectively (Smagghe et al., 2002). However, these compounds did not induce evagination of S. littoralis imaginal discs even at 100 mM or when fed to last instar larvae. In view of the low concentrations of brassinosteroids in plants, any ecdysteroid (ant)agonist activity they may possess will not be biologically relevant. However, given the chemical similarities between brassinosteroids and ecdysteroids, it is of interest to determine which structural modifications are required to convert molecules of one class to others with the biological activity of the other class. Hybrid brassinosteroid/ecdysteroid molecules have been generated and tested in the BII bioassay (for ecdysteroid agonist/ antagonist activity) and in the rice lamina inclination test (for brassinosteroid-like activity) (Voigt et al., 2001). 3.4.6.5. Diacylhydrazines

Owing to the chemical and physical properties of ecdysteroids, it is not possible to regard them as potential insecticidal compounds in their own right

211

(Dinan, 1989). Prospects lie in the identification of nonsteroidal analogs which retain biological activity, but are chemically simpler and more resistant to metabolic and/or environmental degradation. The first nonsteroidal ecdysteroid agonists were identified at Rohm & Haas Co. in the late 1980s (Wing, 1988; Wing et al., 1988). The prototypical molecule was RH-5849 (Figure 6), which was shown to bind to ecdysteroid receptors with relatively low affinity, but, because it is metabolized only slowly, it is able to induce premature molting and death in treated insects. As a result of the synthesis and assessment of a very large number of diacylhydrazines (DAHs), Rohm & Haas could identify and develop analogs as commercially viable insecticides, starting with RH-5992 (tebufenozide) and followed by RH-0345 (halofenozide) and RH-2485 (methoxyfenozide). These compounds possess very low mammalian toxicities and even show some selectivity for certain orders of insects. The DAHs are treated more extensively elsewhere in this series (see Chapter 6.3), and the focus here will be on SARs for DAHs (see Section 3.4.9). There is considerable interest in modeling possible overlaps between DAH and ecdysteroid binding to the ligand-binding domain of the EcR (see below). Although many (165) DAHs have been assessed in the BII bioassay (Table 2), none has yet been identified which possesses antagonistic activity.

Figure 6 Structures of prototypical and commercially relevant diacylhydrazines.

212 Ecdysteroid Agonists and Antagonists

Table 2 Agonistic potencies of diacylhydrazines in the Drosophila melanogaster BII cell assay

RH number

Substituent(s) ring A

Substituent(s) ring B

BII bioassay EC50 (M)

65848 65849 68581 69115 69116 69117 69122 69252 69253 69254 69255 69256 69258 69259 69260 69261 69262 69263 69266 69269 69600 69601 69602 69662 69663 69664 69669 70345 70389 70942 70947 71266 71267 71275 71276 71279 71353 71541 71680 71787 71947 72179 72617 72618 72773 73831 73979 74427 74442 74443 75717 75718 75894

3-Cl H 4-Me 3-OMe 2-NO2 2-Cl 4-CH3 H H H 3-Me 2-Me H H H H H H H H H H H H H H H 4-Cl 2-Cl 3-Cl 2-CH3 H H 4-Me 4-Me 4-Me 4-Cl 4-F H H 4-Ethyl 4-CF3 4-Cl 4-Cl 4-Me H H 4-Et 2-CF3 2-Br 2-NO2 3-CF3 4-n-Pr

3-Cl H 4-Me 3-OMe 2-NO2 2-Cl H 4-Cl 3-Cl 2-Cl 3-Me 2-Me 4-CH3 3-CH3 2-CH3 4-OCH3 3-OCH3 2-OCH3 4-CN 2-NO2 4-F 3-F 2-F 4-CF3 3-CF3 2-CF3 3-CN H H H H 2-Br 4-OH 2-Me 3-Me 2-Cl 2-Me 4-F 4-Phenyl 3-Br H H 3-Cl 4-Me 3-Cl 2-I 4-Acetyl 2-NO2 H H H H 4-Cl

1.8  10
6 1.8  10
6 7.4  10
6 1.2  10
5 5.3  10
6 2.6  10
6 8.3  10
7 1.8  10
5 1.1  10
6 1.3  10
6 2.2  10
6 9.0  10
6 8.0  10
5 2.4  10
6 3.8  10
6 8.2  10
5 1.2  10
5 3.8  10
6 1.2  10
4 1.1  10
6 3.8  10
6 3.3  10
6 6.2  10
6 1.1  10
4 1.1  10
5 3.6  10
6 6.2  10
5 9.0  10
7 8.0  10
6 5.2  10
6 1.1  10
5 8.0  10
7 1.2  10
4 8.0  10
7 6.2  10
7 2.2  10
7 9.3  10
7 4.4  10
6 1.1  10
4 2.1  10
6 8.2  10
7 2.2  10
6 4.2  10
7 1.2  10
5 4.2  10
7 1.2  10
6 1.6  10
5 2.3  10
7 3.5  10
5 3.4  10
5 3.2  10
5 4.3  10
5 9.0  10
6

Table 2 Continued RH number

Substituent(s) ring A

Substituent(s) ring B

BII bioassay EC50 (M)

75916 75992 75995 78295 78374 78404 78434 78435 78447 78448 78449 78487 79186 80664 80665 80678 81204 81215 81252 81398 81539 87590 92895 92897 94496 95028 95075 96586 96596 99547 99613 99633 100153 101292 101570 111016 112608 116715 120004 120064 120230 120436 122632 122659 122972 123238 123757 124863 124948 167174 167179

4-OCH3 4-Et H 2-Cl 3-CH3 2-Cl 4-NO2 4-CN 3-OMe 3-OCH3 3-OMe 2-OMe 4-OH H H 4-F 3-CN S-CH(OH)CH3 3-CO2H 2-F 2-F 2-Et 3-OMe 3-Me 4-I 4-Br 3-Me 3-Cl 4-Acetyl 2-OH 3-OH 3-Cl 2-NH2 2-Me 3-Cl 4-Cl 4-OMe 4-F 3-F 3-F 3-F 3-F 4-F 3-F 4-F 2-F 4-Et 3-OMe 3-OMe 2-NO2 2-NH2

H 3,5-di-Me 3-OH 2-Me H 2-F H H 3-Me H 4-Cl 2-NO2 H 3-NH2 2-NH2 H 3-Me 3-Me 3-Me 3-Me H 3-Me 2-NO2 2-NO2 H 3-Me 3-Cl 4-Me H 3-Me 3-Me 4-F H 2-Cl 2-Cl 2-SMe 3-F 3-Br 2-OMe 2-Me 2-F H 3-OMe 3-OMe 3-F 4-Me 2-OMe 2-OMe 4-F 3-Cl 3-Cl

3.3  10
6 5.3  10
7 2.2  10
4 8.2  10
6 1.3  10
5 1.9  10
5 1.3  10
5 3.6  10
5 2.7  10
6 9.0  10
6 1.6  10
5 3.1  10
5 1.5  10
5 9.3  10
5 2.5  10
5 2.1  10
6 1.8  10
5 5.4  10
6 8.5  10
5 6.1  10
6 5.3  10
6 3.5  10
6 4.0  10
6 1.9  10
6 9.5  10
7 8.5  10
7 2.5  10
6 2.1  10
5 2.3  10
5 1.2  10
5 1.3  10
5 2.3  10
6 4.4  10
6 2.2  10
6 1.2  10
6 2.4  10
6 1.6  10
6 8.7  10
7 2.3  10
6 3.5  10
6 8.0  10
6 4.4  10
6 8.3  10
6 1.1  10
5 1.9  10
6 4.2  10
5 6.0  10
7 1.1  10
5 5.4  10
6 8.3  10
6 1.6  10
6

Test set 75744 75749 75915 78331 78677 78678 79185

4-Et 4-Me 4-OMe 2-Cl 4-CF3 4-CF3 4-i-Pr

3-Et 4-Et 3-Me 3-Cl 4-Cl 3-Me 3-Me

1.5  10
6 9.0  10
5 2.8  10
6 5.4  10
6 7.2  10
6 1.4  10
6 1.7  10
6

Ecdysteroid Agonists and Antagonists

Table 2 Continued RH number

Substituent(s) ring A

Substituent(s) ring B

BII bioassay EC50 (M)

79268 79337 80771 82982 83292 83299 111729 112483 112520 112621 118878 118890

4-CN 4-OH 4-Et 3-OCF3 4-t-Bu 4-t-Bu 4-Et 4-OMe 4-OMe 4-Et 4-F 4-F

3-Me 3-Me 2-F 3-Me 2-NO2 4-F 2-Cl 3-Br 2-Br 3-F 2-OMe 2-NO2

4.2  10
5 4.0  10
5 1.9  10
6 1.4  10
5 3.2  10
6 8.5  10
6 2.2  10
7 8.7  10
7 1.8  10
7 6.0  10
7 1.7  10
6 6.6  10
7 Inactive at 10
4 Inactive at 10
4 Inactive at 10
4 Inactive at 2.5  10
4 >2.5  10
4 Inactive at 10
4 Inactive at 10
3 >2.5  10
5 >1  10
4 >10
4 Inactive at 10
4 >1  10
4 Inactive at 10
4 Inactive at 10
4 >10
4 Inactive at 10
4 Inactive at 10
4 Inactive at 2.5  10
5 Inactive at 10
4 >1  10
3 Inactive at 10
4 Inactive at 10
4 >10
4 Inactive at 10
4 Inactive at 10
4 >1  10
4 Inactive at 10
4 Inactive at 10
4 Inactive at 10
4 Inactive at 10
3 >1  10
3 Inactive at 10
3 Inactive at 10
3

213

now clear that 8-O-acetylharpagide, per se, possesses no agonistic activity and that the activity is attributable to contaminating ecdysteroids (Dinan et al., 2001c). 3.4.6.7. DTBHIB

The synthetic compound 3,5-di-tert-butyl-4-hydroxyN-isobutylbenzamide (DTBHIB) (Figure 7) is reported to be a weak ecdysteroid agonist, as it induces ecdysteroid-specific changes in the D. melanogaster Kc cell line (EC50 ¼ 3  10
6 M) and is able to displace [3H]PoA from receptors in vitro (EC50 ¼ 6  10
6 M). However, the isopropyl analog showed no activity in either test (Mikitani, 1996).

Unquantified weak activity

68583 69118 69123 69267

4-OMe 2-OMe 4-CN H

4-OMe 2-OMe 4-CN 4-NO2

69268 71351 71979 75748 75750 75977 78327 78375 78386 78483 78484 78486 79126 79183

H 4-Cl H H 4-Et 2-Cl 2-F 3-NO2 2-Cl 2-OCH3 2-OMe 2-OMe 4-Et 4-n-Bu

3-NO2 4-Ph 2-OH 4-I 4-Et 4-Et 4-Cl H 4-CN H 3-Me 4-Cl 4-OCF3 4-Cl

80662 80729 80778 80779 81203 83413 89842 96597 112350 119005 126814 141591 141592 141650 141713

4-Et 3-OH 4-CO2Me 4-CO2H 4-Ph 2-Me 4-Et 4-Ac 4-CH3O(CO)4-F 4-Ac 2-Pyrrole H 4-Ethyl 2-Pyrrole

4-CO2Me H 3-Me 3-Me 3-Me 4-Cl 4-OPh 4-Me H 2-OEt 4-Ac 3,5-di-Me 2-Pyrrole 2-Pyrrole H

3.4.6.6. 8-O-Acetylharpagide

The iridoid glucoside 8-O-acetylharpagide (Figure 7), isolated from Ajuga reptans (Labiatae), was reported to possess weak ecdysteroid agonist activity (Elbrecht et al., 1996). Unfortunately, A. reptans is also a rich source of phytoecdysteroids and it is

3.4.6.8. Maocrystal E

As part of an extensive survey of natural products to detect those showing ecdysteroid agonist or antagonist activities in the BII bioassay (Dinan et al., 2001a), it was found that the diterpene maocrystal E (isolated from Isodon spp. (Labiatae)) (Li et al., 1985) showed agonist activity (EC50 ¼ 5 10
6 M). Further, maocrystal E (Figure 7) competed with [3H]PoA for the ligand binding site of D. melanogaster and Choristoneura fumiferana EcR complexes (EcR/USP) with Ki values of 8.6  10
6 M and 5.8 10
5 M, respectively. These data support the interaction of maocrystal E with the ligand-binding domain. 3.4.6.9. Tetrahydroquinolines

Researchers at FMC Corp. (Princeton, NJ, USA) have indicated that 4-phenylamino-1,2,3,4-tetrahydroquinolines also act as ecdysteroid agonists (Dixson et al., 2000). More recently, RheoGene investigators have pursued this chemistry in the context of EcR gene switch actuation (Smith et al., 2003) (see Section 3.4.9.4).

3.4.7. Antagonists 3.4.7.1. Ecdysteroids

Although many ecdysteroids have been identified (Lafont et al., 2002) (see Chapter 3.3), relatively few had until recently been assessed for their biological activity, let alone to determine whether they possess agonistic or antagonistic activity. Apart from ajugalactone (see Section 3.4.3.3), there are no literature reports concerning antagonistic ecdysteroids. Approximately 150 ecdysteroid analogs have now been assessed in the BII bioassay and very recently one of these was shown to possess antagonistic activity. 24x-Hydroxydihydrocarthamosterone (Figure 8) had been isolated as a minor

214 Ecdysteroid Agonists and Antagonists

Figure 7 The structures of 8-O-acetylharpagide, 3,5,-di-tert-butyl-4-hydroxy-N-isobutylbenzamide (DTBHIB), and maocrystal E.

Figure 8 The structure of 24x-hydroxydihydrocarthamosterone.

ecdysteroid from Leuzea carthamoides (Compositae) and has an EC50 value of 2.0  10
5 M versus 5  10
8 M 20E. To verify that the compound is antagonistic, it has been shown that the biological activity co-chromatographs with the ultraviolet (UV)-absorbing peak on high-performance liquid chromatography (HPLC) (Harmatha and Dinan, unpublished data). The stereochemistry of the antagonistic ecdysteroid has not yet been fully elucidated, but it will be interesting to determine what makes this compound antagonistic, while other closely related compounds (e.g., carthamosterone) are agonistic. 3.4.7.2. Brassinosteroids

See Section 3.4.6.4 above. 3.4.7.3. Withanolides

The first evidence that certain withanolides can act as ecdysteroid antagonists was obtained in 1996, when a series of 16 withanolides isolated from Iochroma gesnerioides (syn. I. coccineum: Solanaceae; Alfonso et al., 1991, 1993; Alfonso and Kapetanidis, 1994) were assessed in the BII bioassay (Dinan et al., 1996). None of the compounds showed agonist activity, but several exhibited distinct antagonist activity. The

activities of four of these were adequate for quantification to determine EC50 values (versus 5.0  10
8 M 20E): 2,3-dihydro-3b-methoxywithaferin A (EC50 ¼ 3.5  10
5 M), 2,3-dihydro-3b-methoxywithacnistine (1.0  10
5 M), 2,3-dihydro-3b-methoxyiochromolide (5.0  10
6 M), and 2,3-dihydro3b-hydroxywithacnistine (2.5  10
6 M) (see Figure 9 for corresponding structures) (note that the configuration of the C3 oxygen-containing substituent was determined as 3b after the publication of the paper; Sarker, Sˇik and Dinan, unpublished data). The active compounds possessed an a,b-unsaturated lactone in the side chain, a 3b-oxygen-containing function and a 5b,6b-epoxide. A further 23 purified withasteroids have been assessed for their ecdysteroid agonist/antagonist activities (Dinan et al., 1997a, 2001a). None of these demonstrated antagonistic activity, but one of them, withaperuvin D, was active as an agonist (EC50 ¼ 2.5  10
5 M). This compound also possesses an a,b-unsaturated lactone in the side chain, but the side chain is linked 17a to the ring system. It also possesses a 3a,6a-epoxide with 5b-H, which is unusual amongst withasteroids. Further SAR studies are required to elucidate the structural features required for antagonism and antagonism. 3.4.7.4. Cucurbitacins

Amongst the first 200 plant extracts to be assessed with the BII bioassay for the presence of ecdysteroid (ant)agonists, the methanolic extract of Iberis umbellata (Cruciferae) demonstrated distinct antagonist activity. A literature search revealed that cucurbitacins are characteristic of certain members of this genus (Gmelin, 1963). Bioassay-guided HPLC fractionation of the seed extract gave two UV-absorbing peaks which corresponded to the biological activity. Subsequent identification by nuclear magnetic resonance (NMR) and mass spectrometry revealed them to be cucurbitacin B and cucurbitacin D (Figure 10) (Dinan et al., 1997b).

Ecdysteroid Agonists and Antagonists

215

Figure 9 Structures of withasteroids active as ecdysteroid (ant)agonists.

The purified compounds antagonize the action of 20E (5  10
8 M) in the BII cells; cucB (EC50 ¼ 1.5  10
6 M) and cucD (EC50 ¼ 1.0  10
5 M). Both compounds also compete with [3H]PoA for the ligand binding site of EcR complexes extracted from BII cells, the Kd for cucB being 5.0  10
6 M. CucB also antagonizes the 20E-induced stimulation of a transfected ecdysteroid-regulated reporter gene in D. melanogaster S2 cells and prevents the formation of 20E-induced complex formation of EcR/USP/ EcRE complexes as examined by gel-shift assays. Preliminary SAR studies suggest that the D23-22oxo functional grouping is responsible for the antagonistic activity. In fact, hexanorcucurbitacin D, which lacks this functional grouping, is a weak

agonist and 5-methylhex-3-en-2-one, which mimics C22 to C27 of the side chain of cucurbitacin D, is also a weak antagonist. Thus, cucurbitacins B and D are unequivocal EcR antagonists. Subsequent studies with cucurbitane-type compounds isolated from Hemsleya carnosiflora (Cucurbitaceae) revealed that several of these antagonize 20E action in the BII bioassay (Dinan et al., 1997c). The active principles of further antagonistic plant extracts have been shown to result from the presence of cucurbitacins, e.g., cucurbitacin D from Cercidiphyllum japonicum (Cercidiphyllaceae) (Sarker et al., 1997b) and cucurbitacin D, cucurbitacin F, and 3-epi-cucurbitacin D from Physocarpus opulifolius (Rosaceae) (Sarker et al., 1999b).

216 Ecdysteroid Agonists and Antagonists

Figure 10 Structures of cucurbitacins active as ecdysteroid (ant)agonists.

Figure 11 Structures of limonoids active as ecdysteroid antagonists.

3.4.7.5. Limonoids

The methanolic extract of seeds of Turraea obtusifolia (Meliaceae) showed antagonistic activity. Bioassay-guided purification resulted in the identification of two limonoids, prieurianin and rohitukin (Figure 11) (Sarker et al., 1997a). Neither compound was particularly active (EC50 values ¼ 10
5 M and 1.3  10
4 M, respectively, versus 5  10
8 M 20E). Several other purified limonoids have been assessed for activity (Dinan et al., 2001a), but only nomilin

and obacunanone were found to have weak antagonistic activity. Azadirachtin, which had been suggested to act as an ecdysteroid antagonist, was also inactive (Lehmann et al., 1988; Cle´ ment et al., 1993). It has not yet been determined whether the active limonoids interact with the ligand binding domain of the EcR, somewhere else on the receptor complex or at some other point in the transduction pathway. In view of the wide diversity of natural limonoid structures and their general effects on

Ecdysteroid Agonists and Antagonists

insect feeding, growth, and development (Isman, 1995), a wider range of limonoids should be assessed in the BII bioassay and in a range of ecdysteroid-responsive systems from other insect species to determine the real significance of this class of ecdysteroid antagonist. 3.4.7.6. Stilbenoids

Methanol extracts of seeds of several species of Paeonia (Paeoniaceae) proved to be antagonistic. The antagonistic activity of one of these (P. suffruticosa) was examined more closely and the activity was found to be associated with cis-resveratrol and three oligostilbenes, named as suffruticosols A, B, and C (Sarker et al., 1999a). Further, antagonistic stilbenoids have been isolated from Iris clarkei (Iridaceae) (Keckeis et al., 2000) and Carex pendula (Cyperaceae) (Meng et al., 2001a). The structures of the isolated stilbenoids are presented in Figure 12, while the biological activities are summarized in Table 3. The remarkable observation arising from consideration of the relationship between activity and structure is that the various stilbenoids possess similar activities, even though they differ enormously in molecular size from monostilbenes like cis-resveratrol through di-, tri-, and tetrastilbenes. The stilbenoids from C. pendula (kobophenol B, cis-miyabenol A, and cis-miyabenol C) compete with [3H]PoA for the ligand binding site on the EcR complex (Meng et al., 2001a). It is difficult to envisage how molecules of such differing sizes can interact with the ligand binding domain, although the planar nature of the molecules may facilitate this. 3.4.7.7. Phenylalkanoids

Purified phenylalkanoids (Figure 13) were assessed for ecdysteroid agonist and antagonist activity because they have a wide range of bioactivities, including insecticidal activity (Perrett and Whitfield, 1995). Six phenylalkanoids have been assessed (Dinan et al., 2001a) and all were found to possess antagonistic activity, of which marginatine (EC50 ¼ 5.0  10
5 M versus 5  10
8 M 20E) was found to be the most active. 3.4.7.8. Industrial Compounds

Endocrine disruption is currently an area of considerable, and increasing, concern. Most research to date has been conducted on aquatic vertebrates (fish and mammals) and the emphasis has been placed on disruption involving members of the nuclear receptor superfamily. As EcRs are also members of this superfamily and aquatic arthropods fill important

217

ecological niches and are of great significance in food webs, endocrine disruption in invertebrates could have even more impact, affecting several trophic levels. Research in this area is hampered by restricted knowledge of invertebrate endocrinology (especially with regard to aquatic species) and the vast number of man-made and natural compounds released into the environment, which might potentially act as endocrine-disrupting chemicals (EDCs) (DeFur et al., 1999). The simplicity and robustness of the BII bioassay lend themselves to its use as a preliminary screen to identify potential EDCs, since many compounds can be rapidly tested to determine if they act as ecdysteroid agonists or antagonists. The few active compounds can then be taken forward for more extensive (and very time-consuming) in vivo testing using appropriate invertebrate species. In vivo studies alone very rarely give conclusive information on the mode of action of toxic chemicals, complicating the identification of EDCs by this approach, while a cell-based receptor assay does not reflect the full complications (penetration, metabolism, sequestration, excretion, receptor crosstalk, etc.) of a biological organism. Consequently, the two approaches are complementary and an approach using the BII bioassay as a pre-screen to identify the potentially developmentally disruptive chemicals allows one to proceed to in vivo testing on a rational basis. Several hundred environmental chemicals have now been tested for ecdysteroid (ant)agonist activity in the BII bioassay (Dinan et al., 2001d; Cary et al., unpublished data). Several weak antagonists have been identified, of which diethylphthalate (DEP), bisphenol A (BPA), and lindane (Figure 14) are the most signficant, both in terms of their potencies and the amounts released into the environment (Dinan et al., 2001d). For each of these three compounds we could demonstrate that the antagonistic activity co-chromatographs with the named compound (i.e., is not associated with an impurity) and that the compound competes with [3H]PoA for the ligand binding domain of the EcR complex. However, the EC50 values (2.0  10
3 M, 1.0  10
4 M, and 3.0  10
5 M for DEP, BPA, and lindane, respectively; versus 5  10
8 M 20E) are high and, given the concentrations of these compounds in the environment, it is unlikely that they are true invertebrate EDCs, although the caveats about the differences between in vitro and in vivo assessments mentioned above and the possibility that other species could be more susceptible than D. melanogaster should be borne in mind. Others are also beginning to use in vitro assays for the identification of potential invertebrate EDCs (Oberdo¨ rster et al., 1999, 2001).

218 Ecdysteroid Agonists and Antagonists

Figure 12 Structures of stilbenoids active as ecdysteroid antagonists.

Ecdysteroid Agonists and Antagonists

219

Table 3 Antagonistic activities of stilbenoids in the Drosophila melanogaster BII cell bioassay (vs. 5  10
8 M 20-hydroxyecdysone) Number

Compound

Source

EC50 (M)

Monostilbenes

cis-Resveratrol trans-Resveratrol

Paeonia suffruticosa

Distilbenes Tristilbenes

Ampelopsin B cis-Miyabenol C Suffruticosol A Suffruticosol B Suffruticosol C a-Viniferin Kobophenol B cis-Miyabenol A

Iris clarkei Carex pendula Paeonia suffruticosa Paeonia suffruticosa Paeonia suffruticosa Iris clarkei Carex pendula Carex pendula

1.2  10
5 Inactive 3.3  10
5 1.9  10
5 5.3  10
5 1.4  10
5 2.2  10
5 1.0  10
5 3.7  10
5 3.1  10
5

Tetrastilbenes

Figure 13 Structures of phenylalkanoids active as ecdysteroid antagonists.

3.4.8. Functions of Natural Ecdysteroid (Ant)Agonists in Insect–Plant Relationships 3.4.8.1. Agonists

When phytoecdysteroids were first discovered, it was suggested that they were a defense against insect predators (Galbraith and Horn, 1966). However, it soon became clear that certain insect species are highly resistant to ingested ecdysteroids (review: Dinan, 1998). It is now generally accepted that phytoecdysteroids contribute to deterrence of nonadapted phytophagous invertebrate predators, either by acting as antifeedants (mediated by taste receptors) or as hormonal toxicants on ingestion. Some 5–6% of terrestrial plant species accumulate phytoecdysteroids (Imai et al., 1969), of which approximately half contain the compounds at levels which in themselves would deter nonadapted insect

Sigma

predators (Dinan, 1995a). Ecdysteroids may still contribute to the protection of plant species containing lower levels, since the ecdysteroids may interact synergistically with other secondary compounds. There appears to be a relationship between the occurrence and levels of phytoecdysteroids in the food plants of a particular insect species and its ability to cope with ecdysteroid in its diet (Blackford and Dinan, 1997). Thus, monophagous species such as larvae of the peacock butterfly (Inachis io) which feeds on ecdysteroid-negative stinging nettle (Urtica dioica; Urticaceae) would rather starve to death than consume nettle leaves coated with even low levels of 20E, while larvae of the oligophagous cinnabar moth (Tyria jacobeae) which encounter some ecdysteroid-containing plants in its host-range are moderately resistant to ingested ecdysteroids, but suffer developmental defects on the ingestion of larger amounts. No studies have been yet performed on the role of maocrystal E in insect–plant interactions. 3.4.8.2. Antagonists

It has been realized for some time that compounds such as cucurbitacins, withanolides, stilbenoids, phenylalkanoids, and limonoids contribute to plants’ protection strategies against invertebrate predators (Ascher et al., 1980; Govindachari et al., 1995; Miro´ , 1995; Perrett and Whitfield, 1995), but little was known about the mechanism(s) of action. The characterization of members of these classes of compounds as EcR antagonists has provided a testable hypothesis for the investigation of their roles in insect–plant relationships. All the active compounds identified so far have relatively weak affinities for the D. melanogaster EcR complex (Kd values in the mM range). This may reflect the low cost to the plant in producing these compounds and the high energy cost to phytophagous insects of detoxifying large amounts of such compounds (Dinan et al., 2001b).

220 Ecdysteroid Agonists and Antagonists

Figure 14 Structures of potential environmental disruptors interacting with ecdysteroid receptors.

3.4.9. QSAR and Molecular Modeling

Table 4 Ecdysteroid structural modifications and contributions to BII p(EC50)

3.4.9.1. Ecdysteroid SAR and QSAR

The early literature on ecdysteroid SAR studies has been reviewed (Bergamasco and Horn, 1980; Horn and Bergamasco, 1985; Dinan, 1989). The various bioassays employed vary in their complexity of performance and interpretation. A number of factors can affect the potency of a test compound to a greater or lesser extent depending on the assay system: penetration, metabolism, excretion rate, sequestration, and target site activity. In vivo assays will give a measure of all of these acting in concert, but not allow the individual contributions to be identified. On the other hand, cell-free receptor assays probably give a much more direct measure of target activity, but may bear little resemblance to the in vivo situation. Horn and Bergamasco (1985) summarized their main conclusions regarding ecdysteroid SAR in insect systems as follows: . . . . .

an A/B cis-ring junction is essential a 6-oxo-7-ene grouping is essential a full sterol side chain is essential a free 14a-hydroxyl group is essential 2b-, 3b-, and 20R-hydroxyl groups enhance biological activity . a C25 hydroxyl diminishes activity. These conclusions were empirically derived from data combined from a number of different bioassay systems (mainly the Calliphora, Musca, and Sarcophaga assays, the Chilo dipping test and the D. melanogaster Kc cell assay). On the basis of the above summary, Bergamasco and Horn (1980) visualized the ecdysteroid interacting with the ligand binding pocket of the receptor at three major sites: the bface of the A- and B-rings, the a-face in the region of the 14a-hydroxyl group, and the side chain from C22 to C27. Later, uniform data from the BII bioassay enabled quantification of the potency contributions to the BII EC50 by individual structural features, by making direct pairwise comparisons between structures which differ only in a single feature (Table 4) (Dinan et al., 1999c).

Potency-enhancing modifications

Dp(EC50)

þ2b-OH þ20-OH þ14a-OH þ22R-OH

2.28 1.35 0.60 0.47

22-OH ! 22-›O

0.04

Potencydiminishing modifications

3b-OH ! 3-›O b-C-3 ! a-C-3 þ5b-OH trans ! cis C/D-junction 6-›O ! 6-OH (a or b) þ11b-OH 20R-C- ! 20S-C 22R-OH ! 22S-OH þ22-OH, þ25-OH þ24-CH3 (R or S ) þ25-OH 25-OH ! 24/25 C›C

Dp(EC50)


0.74
0.0
0.25
1.44
1.89
1.87
1.72
0.60
0.34
0.85
0.89
0.91

In this way the previous SAR could be qualitatively substantiated and enhanced. Significantly, the previous SAR formulation described primarily as a brief collection of essential molecular features could now be cast in terms of the cumulative contribution, both enhancing or diminishing, of an expanded feature set (Table 4). In the language of pharmacophore analysis, the most highly contributing features are hydrogen bond donors or acceptors attached to C2, C3, C6, C20, and C22 together with a lipophilic group extending from C22. These groups are arranged approximately according to the geometry shown in Figure 15, wherein the ring sizes and ring junction stereochemistry determine the exact distances and angles. No single functional group is essential for ecdysteroidal potency; noteworthy is that the 14a-OH group can be replaced with unsaturation or can be altogether removed without abolishing BII bioassay activity. Available data conflict as to a uniformly positive contribution of the 14a-OH to the pharmacophore. More recent work involving examination of selected ecdysteroid probe sets in engineered EcR mammalian gene switch assays or in Sf-9 cells

Ecdysteroid Agonists and Antagonists

221

Figure 15 Ecdysteroid pharmacophore. Features are generalized where possible; HA, hydrogen bond acceptor; HD, hydrogen bond donor; (
), polar negative atom.

is roughly consistent with the previous SAR (Nakagawa et al., 2000a; Saez et al., 2000). The available ecdysteroid datasets are unfortunately too limited for quantitative analysis by classical QSAR. However, quantitative analysis was performed on the BII data set using the multidimensional comparative molecular field analysis (CoMFA) approach (Dinan et al., 1999c; Ravi et al., 2001). By the CoMFA method, the intensity of steric and electrostatic fields at specified points in a threedimensional lattice is measured and tallied as a function of each member ecdysteroid of the QSAR training set. Thereby, a descriptor table of many columns is generated. Partial least-squares supplies the appropriate statistical treatment to manipulate the data and derive a QSAR expression. This expression is best depicted as color-coded surfaces or polyhedra describing regions in space where the presence of generalized pharmacophoric features would be expected to either enhance or diminish efficacy. For the ecdysteroids, a CoMFA model was developed with an r2 ¼ 0.92 and a cross-validated r2 (q2) of 0.593 for a 67-member training set and an average residual of fit of 1.47 p(EC50) units for a 16-member test set (Figure 16). The model is characterized by a large (þ)-charge-enhancing region (blue) wrapped around the steroid side chain and (þ)-charge-diminishing regions (red) at O20, between C26 and C27, and below C7/C8. Stericenhancing regions (green) appear distal to C21 and at the terminus of the side chain. Steric-diminishing regions (yellow) appear around the perimeter of the side chain. The CoMFA is consistent with, and extends, the simple functional group-oriented pharmacophore of Figure 15, and provides an abstraction useful for evaluating new or hypothetical structures which might not fit into the simple functional group schema.

Figure 16 Comparative molecular field analysis (CoMFA) electrostatic field contour plot (standard deviation  coefficient) for favored ecdysteroid CoMFA model. Blue/red polyhedra represent regions where positive charge is favored/disfavored; contribution level ¼ 85/8%, respectively. Green/yellow polyhedra represent sterically favored/disfavored regions; contribution level ¼ 80/25%, respectively. Ponasterone A (PoA) is depicted. (a) View from the b-face; (b) view towards the C6/ C14 edge of the steroid. (Reproduced with permission from Ravi M., Hopfinger A.J., Hormann, R.E., Dinan, L., 2001. 4D-QSAR analysis of a set of ecdysteroids and a comparison to CoMFA modeling. J. Chem. Inform. Comput. Sci. 41, 1587–1604; ß American Chemical Society.)

The four-dimensional QSAR (4D-QSAR) method (Hopfinger et al., 1997) provides another complementary view of ecdysteroid QSAR in the BII assay. In 4D-QSAR, structures for analysis are passed through a molecular dynamics simulation, and the QSAR descriptors are derived from the frequency that particular atom types might populate individual cells in a three-dimensional lattice enveloping the training set structures. Thus, in contrast to CoMFA, 4D-QSAR takes conformation into account. The method can help discriminate among ligand alignment postulates and can provide optimal ligand conformation hypotheses. Data reduction is accomplished by partial least-squares, and models are selected and optimized by use of a genetic algorithm. 4D-QSAR was applied to essentially the same training set as used for the CoMFA modeling, resulting in four complementary models (r2 ¼ 0.83–0.88, q2 ¼ 0.76–0.80), one of which is depicted in Figure 17 (Ravi et al., 2001). Taken together, the 4D-QSAR

222 Ecdysteroid Agonists and Antagonists

Figure 17 One of the favored ecdysteroid four-dimensional quantitative structure–activity relationship (4D-QSAR) models relative to PoA in its postulated active conformation. Alignment is based on atoms 6, 12, and 16. No external whole molecule descriptors were used in this model. The grid cell occupancy descriptors (GCODs) are shown as spheres. Interactive pharmacore element (IPE) type abbreviations: ANY, any atom (green/yellow); NP, nonpolar (white/gray); (
), polar negative (red/blue); HD, hydrogen bond donor (cyan/pink); HA, hydrogen bond acceptor (pink/cyan). Positive values of a GCOD indicate that occupancy by the appropriate IPE increases activity while a negative value indicates occupancy decreases activity. (Reproduced with modification and permission from Ravi M., Hopfinger A.J., Hormann R.E., Dinan L., 2001. 4D-QSAR analysis of a set of ecdysteroids and a comparison to CoMFA modeling. J. Chem. Inform. Comput. Sci. 41, 1587–1604. ß American Chemical Society.)

p0240

models corroborate the CoMFA conclusions, and also specify structural requirements more precisely. The 4D-QSAR models indicate that the hydroxyl group at C2 should be a hydrogen bond acceptor, the hydroxyl at C20 should be a hydrogen bond donor, the atom attached to C22 should be polar and bear a partial negative charge (and is a likely hydrogen-bond acceptor, although HA/HD role swapping with O20 is possible), and that there should be no hydrogen bond acceptor at C25. The general picture generated by the ecdysteroid CoMFA and 4D-QSAR models is that in the ligand binding cavity of the EcR, the steroid side chain lies in a sterically restrictive hydrophobic cylinder with hydrogen-bonding functionality near O20 and O22. Interestingly, analysis of weakly active ecdysteroids in the 4D-QSAR models indicates that the potency depression is often due not so much to an offending functional group as it is to the inability to assume a non-offending conformation. Homology modeling of the EcR ligand-binding domain and ecdysteroid docking generated several interesting hypotheses for ecdysteroid–EcR binding, notwithstanding serious challenges owing to the absence of homologous nuclear receptors with identity any greater than ca. 30%, and, of course, the absence of a publicly available EcR crystal structure itself (see Chapter 3.5). Wurtz et al. (2000) have

proposed two models for ecdysteroid interaction with the dipteran C. tentans EcR ligand binding domain, based on homology models built from the ligand binding domains of the human retinoic acid receptor (hsRARg) and the human vitamin D receptor (hsVDR). Both models insert the steroid side chain in a ‘‘tail-first’’ orientation, such that ring-A remains closest to the enclosing Helix-12. Between these two models, the orientation of 20E differs by roughly 180 , thereby altering the interactions between the functional groups on 20E and the residues lining the ligand binding pocket. Another model built at Rohm & Haas, based on estrogen receptor (ER), thyroid harmone receptor (TR), and vitamin D receptor (VDR) and docked with 20E in the ‘‘tail-first’’ orientation, was successfully used to guide mutant generation for selective modulation of ecdysteroid responsiveness, while leaving DAH sensitivity unaltered (Kumar et al., 2002). A more recent model has been constructed by investigators at Sankyo and Nippon Kayaku (Kasuya et al., 2003). The CoMFA and 4D-QSAR view of a hydrophobic cylinder surrounding the side chain, the hydrogen bond acceptor characteristics of 22OH, and the hydrogen bond donor characteristic of 20OH can all be easily reconciled with the CtEcR (C. tetans) 20E-liganded homology models based on either the retinoic acid receptor (RAR) or the VDR, as well

Ecdysteroid Agonists and Antagonists

p9000

as with the generally hydrophobic ligand binding domain characteristics of known nuclear-receptor ligand binding domains on which these models are based (Wurtz et al., 2000). On the other hand, the CtEcR homology models predict that all six hydroxyls and the C6 carbonyl of 20E are implicated in H-bonding, whereas the receptor-independent 4D-QSAR and CoMFA models do not recognize 14OH in any significant way, and strongly suggest that 25OH is not involved in hydrogen bonding. A fortiori, simple empirical SAR of the 25 position indicates that 25OH is detrimental to EcR affinity. Despite these conflicts, the bulk of empirical SAR, multidimensional QSAR analysis, homology modeling, and docking generally provide a consistent picture of the factors involved in ecdysteroid activity. While it is significant and reassuring that the calculational models and qualitative SAR are generally in concordance with each other and with ecdysteroid binding data and physiological data, a more stringent test is in consonance with a crystal structure of EcR bound with an ecdysteroid. The ligandbinding domains of EcR/USP from Heliothis virescens (HvEcR) co-crystallized with PoA at 3 A˚ resolution considerably enlightens our understanding of ecdysteroid SAR (Billas et al., 2003). In this structure, PoA assumes a pose similar to vitamin D in VDR, in which the steroid assumes a crescent shape with its side-chain proximal to Helix-12 (‘‘headfirst’’), and the A-ring is held in a chair conformation. Hydoxyl groups at carbons 2 (R383), 3 (E309), 14 (T343/T346), and 20 (Y408) as well as the C-6 carbonyl (A398) all participate in hydrogenbonding. Moreover, the interacting residues are generally conserved across insect species, which concurs with the ubiquity of PoA recognition. The HvEcR residues near the side-chain terminus are generally hydrophobic, consistent with qualitative SAR and QSAR models, which all discount 25-OH as a pharmacophore element. The combined CoMFA and 4D-QSAR models correctly interpreted/predicted the crescent shape, A-ring chair conformation, hydrophobic quality, and steric permissiveness of the distal portion of the side-chain, and the identity of most hydrogen-bonding units (Figure 15). However, whereas 4D-QSAR models predicted that C-22 would be involved in hydrogen bonding as an acceptor, the HvEcR crystal structure indicates no such role. Likewise, although qualitative SAR and QSAR models lead one to the conclusion that the C-14 hydroxyl may not necessarily be advantageous for binding, the HvEcR crystal structure clearly shows not one, but two hydrogen-bonding interactions at this position.

223

In interpreting the immensely valuable EcR crystal structures, one should keep in mind that dynamics exert a significant role in ligand–protein interactions, and that multiple ligand-binding modes may be possible. For example, some of the homology modeling that antedates the HvEcR–PoA crystal structure hypothesizes a ‘‘tail-first’’ ecdysteroid orientation – a hypothesis testable through careful docking studies and possibly multiple crystal structures. Although hydrogen-bonding HvEcR residues are generally conserved across species, nonconserved binding pocket residues may provide a means for supplementary or substitute binding interactions, particularly if the specific ecdysteroid analog in question offers complementary features for such hypothetical alternative interactions. 3.4.9.2. Diacylhydrazine SAR and QSAR

3.4.9.2.1. Lepidopterans The diacylhydrazines (Figure 18) have enjoyed the most extensive qualitative and quantitative analysis, owing to commercial interest and the relative abundance of analogs. The chemotype was first synthesized and identified as an insecticide in the early 1980s (Hsu and Aller, 1991), and shortly thereafter reported as an ecdysteriod agonist (Wing, 1988; Wing et al., 1988). Using Kc cells, competitive binding studies indicated that RH-5849 (1,2-dibenzoyl-1-tert-butylhydrazine) is a competitive inhibitor with PoA with a Kd value of 2 mM, which implies that the two ligands share a common binding domain. Depression of the PoA kinetic on-rate, but not the off-rate, by RH-5849 are consistent with affinity for a common steroid/ DAH binding site rather than an allosteric site. DAH toxicity to caterpillars is attributable to hyperecdysteroidism and premature initiation of molting. For an initial set of 28 DAHs, Kc cellular response is proportional to propensity toward head capsule apolysis in the tobacco hornworm, Manduca sexta (Wing et al., 1988). Hsu (1991) described an initial

Figure 18 Diacylhydrazine (DAH) notation and numbering system.

p9005

224 Ecdysteroid Agonists and Antagonists

DAH SAR for toxicity toward southern armyworm (Spodoptera eridania) as follows: 1. A- and B-regions: when A or B is phenyl and C is tert-butyl, toxicity is retained when the alternate A or B group is an aryl, aromatic heterocycle, aliphatic cycle or short alkenyl. Acyclic alkyl or benzyl are somewhat less active. 2. C-region: a bulky group is essential, but excessive size significantly truncates potency. Tertbutyl confers the optimum blend of foliar and plant systemic potency. Some functionality, such as cyano, is permissible. Later, Rohm & Haas investigators updated the empirical SAR for southern armyworm toxicity (Hsu et al., 1997): 1. A- and B-region: aryl > heterocyclic; asymmetric A/B-substitution > symmetric A/B-substitution. 2. A-region (monosubstituted, B ¼ phenyl): 2-Cl, 2-CH3, 3-Cl, 3-OCH3, 4-Et, 4-Br, 4-I, 4-CH3, 4CF3, 4-OCH3, > H > 2-Br, 2-NO2, 2-CF3, 2-NH2, 2-OCH3, 3-CH3, 3-Et, 3-NO2, 4-Cl, 4-iPr, 4-t-Bu. 3. B-region (monosubstituted, A ¼ phenyl): 2-ethyl, 2-I, 2-Br, 2-NO2, 2-Cl, 3-Br, 3-ethyl, 3-methyl, 3-Cl, 4-Cl, 4-F > H > 2-OCH3, 2-CH3, 2-NH2, 3CF3, 3-OCH3, 3-NO2, 3-NH2, 4-NO2, 4-CH3, 4-OCH3. 4. D-region: H or hydrolyzable group. 5. Backbone: C ¼ O > C ¼ S, S ¼ O; NH–N(tbu) > CH2–N(tbu), NH–CH(tbu). Significantly, nonlinear, intra-ring, and interring substituent relationships were observed, thereby presenting a challenge for lead optimization which nonetheless proceeded steadily to the invention of RH-5992 (tebufenozide), and later advanced to the development of RH-2485 (methoxyfenozide), insecticides. More recent reports from Sankyo and Nippon Kayaku Co. describe qualitative DAH SAR pertaining to structures related to chromafenozide insecticide (Sawada et al., 2003a, 2003b, 2003c). Various DAHs with heterocycles fused to the 3,4-positions of the A-ring are toxic toward the common cutworm (Spodoptera litura). The most active compounds have oxygen and carbon-containing, five- or sixmembered fused rings devoid of bulky or hydrogen bond-donating substitution. Optimized structures also bear a small substituent at the A2 position, typically methyl (a pattern shared with methoxyfenozide), while the remainder of the molecule reflects the structure of RH-5992. With respect to the C-module, considerable potency is retained when neopentyl, 1-t-butyl-ethyl, or 1,1-dimethylbenzyl replace the t-butyl group. Hence, once the

A- and B-modules are optimized, the C-module becomes less refractory towards structural modulation. Many of these observations are implicit in earlier DAH patent literature; however, patent data format generally does not lend itself to SAR analysis. Additional D-module SAR from Sankyo and Nippon Kayaku (Sawada et al., 2003c) and the observation that dibenzoyl cyclic hydrazides are inactive (Toya et al., 2002) remain consistent with the earlier hypothesis that only hydrolyzable or readily metabolizable groups are tolerated in the D-module, and that the bioactive chemical species at the EcR is a DAH with a free amide NH. Notwithstanding, the observation that alkoxycarbonylmethyl groups are tolerated as the D-module does challenge the earlier, more restrictive hypotheses concerning the amide NH (Cao et al., 2001). The DAH chemotype is well suited for Hansch– Fujita QSAR (Hansch and Leo, 1995), and a number of QSAR relationships have been proposed by Nakagawa and colleagues to describe aspects of DAH insect toxicity under varying circumstances. The Hansch approach considers combinations of substituent physiochemical properties and other simple molecular properties as descriptors of the biological activity. The result is a multiple linear regression equation, which, if demonstrated to be robust statistically, can be quite instructive in its quantitative description of the structural factors responsible for potency. In examination of DAH toxicity to larvae of C. suppressalis for a series of 46 DAHs unsubstituted on the A-ring and monosubstitued on the B-ring, it was determined that the level of toxicity is positively influenced by elevated log P and elevated electronic inductive effect (sI) of the ortho-substituent (Oikawa et al., 1994a). Potency is suppressed by voluminous meta- and para-substituents as well as multiple (2, n) substitution patterns. The same training set with Chilo data was later examined by simply measuring the distance from the A-carbonyl oxygen to the attachment atom of the B-subsitutent for DAHs represented in a crystal structure-like conformation (Qian, 1996). A parabolic relationship was found with an optimal distance of ca. 4.6 A˚ . An almost identical training set was used to determine the SAR for toxicity toward larvae of the beet armyworm (Spodoptera exigua); this turned out to be highly correlated with toxicity toward Chilo larvae, and could be described by a very similar regression equation (n ¼ 41; s ¼ 0.275; r2 ¼ 0.959; descriptors: log P, sI, Vortho, meta, para, indicator variables for mutliply substituted members). For a complementary training set in which the B-ring was held constant with a near-optimized 2-Cl substituent and

Ecdysteroid Agonists and Antagonists

the A-ring pattern was varied, the following equation for Chilo toxicity was derived: pLD50 ¼ ð0:722ÞDlog P
ð0:74ÞSDLortho
ð0:868ÞSDVwmeta
ð0:485ÞDLpara þ 6:653 n ¼ 44; s ¼ 0:284; r ¼ 0:896; F4;39 ¼ 39:5

½1

This equation states that for the training set with A-ring variation, toxicity to Chilo larvae is positively influenced by high log P and negatively influenced by the length of substituents in the ortho- and parapositions and the summed volume of the metasubstitutents (Oikawa et al., 1994b). Thus, the most active DAHs in this series bear small lipophilic substituents which increase the log P, and therefore pLD50, to a greater degree than their size detracts from activity. When a very similar training set was tested against beet armyworm larvae, an exactly analogous regression equation was developed (n ¼ 42, s ¼ 0.273, r2 ¼ 0.902, descriptors: log P; DLortho ; Vwmeta ; Lpara ) reflecting toxicity highly correlated with that of Chilo (Smagghe et al., 1999). Several related QSAR studies were perfomed with Chilo in which the ecdysteroid-agonist response was measured not by toxicity, but rather by the rates of chitin synthesis in cultured integument fragments from diapause larvae (Nakagawa et al., 1995b, 2000a). The first study on 37 B-2-Cl and A-unsubstituted DAHs, members of the training sets from the two prior studies on Chilo larvae, also resulted in a linear regression with terms resembling the sum of terms from the two prior Chilo studies (n ¼ 37, s ¼ 0.288, r2 ¼ 0.872, F9,27 ¼ 20.43). The second study used a training set consisting of 23 DAHs heavily represented by structures in which one or the other aryl ring was replaced by a saturated ring or chain. Here, a QSAR was developed which described integument activity in terms of a positive influence by log P and a negative influence by the distances between the carbonyl carbons and the distal carbon atoms on the same side of the hydrazine linkage. In a Hansch study using larval toxicity of the spruce budworm, a set of 10 A- or B-ring monosubstituted DAHs showed a positive correlation of toxicity to substituent p-lipophilicty values and a negative correlation to substituent length (L), more or less independent of substituent position (Mohammed-Ali et al., 1995). The classical QSAR approach is attractive for its simplicity. It can provide straightforward insights for ligand design. The method is limited, however, by application to datasets with a common core region, and classical QSAR’s potential for underparameterization can easily lead to oversimplification (Hansch and Fujita, 1995). Addressing these issues

225

in the context of DAH QSAR, the three-dimensional CoMFA approach has been applied to a balanced set of 37 A- and B-ring substituted DAHs aligned in the crystal structure conformation, and using ecdysteroidal-responsive N-acetylglucosamine incorporation of Chilo integument as the dependent variable. The steric field exerts the most dominant role in the model (52.5% contribution) with potency-diminishing contours located in A- and B-ring positions reassuringly consistent with the two Chilo Hansch analyses described above (r2 ¼ 0.845, q2 ¼ 0.438, s ¼ 0.292, four components). log P contributes to a level of 10.7%; the remainder of the descriptors arise from the electrostatic field (Nakagawa et al., 1995a). 3.4.9.2.2. Nonlepidopterans The initial report of the ecdysteroidal properties of DAHs correlated the hyperecdysonistic response in larvae of a lepidopteran, M. sexta with the affinity for a dipteran, D. melanogaster EcR in Kc cells for an early set of DAHs (Wing, 1988; Wing et al., 1988). This observation presaged research in defining the physiological and physiochemical reasons for the exquisite lepidopteran selectivity of DAHs. The general understanding which has emerged is that: (1) species susceptibility to DAH toxicity is primarily a function of the intrinsic affinity of the DAH to the specific EcR, (2) the general DAH chemotype has the greatest affinity for lepidopteran EcRs; DAH subsets have moderate affinity for coleopteran and dipteran EcRs, (3) ecdysteroid affinity is fairly consistent across insect species while DAH affinity modulates above or below 20E or PoA potency levels, depending upon species, (4) absorption, distribution, metabolism, excretion, and insect behavior may modulate, but do not usually significantly affect, the toxicity levels determined primarily by EcR affinity (Smagghe et al., 2001). These general conclusions have been thoroughly reviewed (Hsu et al., 1997; Dhadialla et al., 1998; Carlson, 2000). The isolation of the locus of biological activity and selectivity to the EcR enables the interpretation of QSARs in terms of receptor binding pocket interactions and facilitates direct quantitative comparisons across species. For the coleopteran Colorado potato beetle (Leptinotarsa decemlineata), fourth instar larvae were monitored for toxicity after topical application of a set of 45 A-ring unsubstitued DAHs (Nakagawa et al., 2001b). Unlike a closely analogous DAH set examined in C. suppressalis, which showed a uniformly augmenting effect ) and a negative influence by from log P and (sortho I Vwmeta and multiple substitution, DAH toxicity in L. decemlineata exhibits a more complex convex

226 Ecdysteroid Agonists and Antagonists

parabolic relationship to hydrophobicity, which can be expressed in terms of the p-value of substituents in the meta-position, and which is enhanced by large ). Only the diminortho-substitents (Vwortho ; DBortho 1 ishing effect of large para-substitents (C. suppressalis: Vwpara , L. decemlineata: DBpara 5 ) is analogous across these two species. Overall, there is no clear correlation of toxicity between these two species. Interestingly, the ortho-sec-butyl substituent was found to be quite favorable for activity against L. decemlineata. Examining the substituent effects on the A-ring while the B-ring is held constant as 2-Cl (another DAH set closely analogous to one described above for C. suppressalis) the following equations were developed to describe two distinct DAH subclasses (Nakagawa et al., 1999): pLD50 ¼ ð0:887Þlog P þ ð0:395ÞDBortho 5 þ ð0:435ÞEmeta þ ð0:65ÞEpara þ ð1:09ÞHB þ 2:97 s s

n ¼ 28; s ¼ 0:328; r2 ¼ 0:842; F5; 22 ¼ 10:68

½2

pLD50 ¼ ð
1:695Þlog P
ð0:527ÞSEortho s
ð0:766ÞSEmeta
ð0:916ÞSEpara þ 9:23 s s n ¼ 16; s ¼ 0:373; r2 ¼ 0:885; F4;11 ¼ 9:982

½3

Here, the effect of hydrophobicity is expressed in two equations with a single log P term rather than a multiterm expression in one equation. One DAH subclass, which is more populous and tends to be monosubstituted, increases in toxicity with increasing log P (eqn [2]), while the second subclass, generally multisubstituted, shows the opposite effect (eqn [3]). As with C. suppressalis toxicity, in L. decemlineata, the first subclass exhibits a toxicity-intensifying effect from log P, but unlike C. suppressalis, a diminishing effect from sterically requiring substituents in all positions (cf. eqn [1]). For the second subclass, the situation is completely reversed with respect to log P and substituent sterics. To highlight the dichotomy of lepidopteran versus coleopteran SAR, the lepidopteran-optimized DAHs, tebufenozide and methoxyfenozide, were observed to be at least two orders of magnitude weaker than the coleopteran-optimized DAH, halofenozide, in L. decemlineata (pLD50 ¼ 3.38, 3.29 versus 6.09), while in the lepidopteran S. exigua, almost the opposite is the case (pLD50 ¼ 7.5, 8.18 versus 6.1; Smagghe et al., 2003). A direct comparison of L. decemlineata toxicity with that of C. suppressalis and S. exigua by the CoMFA method illuminates the disparate balance of structural factors responsible for toxicity in the larval stages of these organisms (Smagghe et al., 2003). Earlier toxicity data from all three organisms

for a common set of 59 DAHs exemplifying modifications on both rings was used to build CoMFA/ CoMSIA (comparative molecular similarity indices analysis) models (Klebe, 1998) by a uniform procedure. All models retained a significant steric component. However, whereas the resultant S. exigua and C. suppressalis models greatly benefited from hydrophobic field and surface area contributions, as well as other factors, the favored L. decemlineata model was dominated by sterics with an additional contribution only from electrostatics (for all three models, r2 ¼ 0.843–0.958, q2 ¼ 0.583–0.683, 5–6 components). The models were optimized on test set r2 (24–50 test set members) as well as training set q2 in order to produce a QSAR tool which would be predictive as well as explanatory (adjusted test set r2 ¼ 0.40–0.465). Another CoMFA analysis was performed on a balanced set of 100 DAHs modeled in the folded crystal structure conformation (Chan et al., 1990) and tested in the Drosophila BII assay (Dinan, 2003; Dinan and Hormann, unpublished data). Many of the training set members examined against C. suppressalis and L. decemlineata appear in the Drosophila model. Sterics, electrostatic, and hydrogen bond donor/acceptor fields all play a balanced role with log P in explaining ecdysonergic activity (r2 ¼ 0.903, q2 ¼ 0.737, five components). Not only do the field contributions differ from prior lepidopteran or coleopteran models, the Drosophila BII field contours vary significantly as well (Figures 19 and 20). In addition to the native inducible gene expression systems discussed so far, engineered heterologous systems may shed light not only on ligand–EcR recognition, but also on secondary factors which could modulate the specific EcR–ligand interaction. A set of 146 DAHs spanning considerable substituent space were analyzed in a gene switch derived from Bombyx EcR and utilizing endogenous RXR as partner protein and b-galactosidase as reporter in stably transfected human kidney cells (Suhr et al., 1998; Hormann et al., 2002). The ligands were modeled in the folded conformation. From these data, CoMFA/CoMSIA (r2 ¼ 0.633, q2 ¼ 0.831, five components) and 4D-QSAR models (r2 ¼ 0.709, q2 ¼ 0.644) were constructed and evaluated with a 30 member test set (average residual fit ¼ 0.493– 0.521 pEC50 units). The favored CoMFA models utilized hydrogen bonding fields and the CoMSIA hydrophobic field in addition to steric and electrostatics in rather complex contours. A large hydrophobic contour appears in the region between the two aromatic rings. The favored 4D-QSAR model is characterized by several aromatic grid cell occupancy descriptors in addition to the more common

Ecdysteroid Agonists and Antagonists

227

Figure 19 CoMFA electrostatic and steric field contour plot (standard deviation  coefficient) for DAH CoMFA model of BII data. Blue/red polyhedra represent regions where positive charge is favored/disfavored; contribution level ¼ 75/25%, respectively. Green/yellow polyhedra represent sterically favored/disfavored regions; contribution level ¼ 65/35%, respectively. Tebufenozide is depicted.

Figure 20 CoMFA hydrogen bond field contour plot (standard deviation  coefficient) for DAH CoMFA model of BII data. Blue/red polyhedra represent regions where hydrogen bond donation is favored/disfavored; contribution level ¼ 70/30%, respectively. Green/ yellow polyhedra represent hydrogen bond acceptor favored/disfavored regions; contribution level ¼ 70/30%, respectively. Tebufenozide is depicted.

any-atom and nonpolar atom type descriptors; most grid cell occupancy descriptors (GCODs) are clustered around the more distal regions of the aromatic rings. Model building with a number of trial alignment hypothesis strongly suggests that the DAHs align in the Bombyx mori EcR cavity principally according to one or more of the backbone atoms, in contradistinction to one or the other of the benzamide fragments. 3.4.9.2.3. Pharmacophores The pharmacophoric features common to all sensitive insect orders are: 1. Two hydrogen acceptor or polar negative atoms space approximately approximately 3.5–4.0 A˚ apart. 2. A bulky, conformationally-determining (see below) lipophilic group located asymmetrically between the two negative centers.

3. Moderately sized (about six carbons) groups on either side of the negative centers. 4. A hydrogen bond-donating group is highly advantageous when located near the alternate negative center. The pharmacophore has been most aptly manifested in mono-N-substituted diacylhydrazides. Toxicity optimized for both lepidopterans and coleopterans favors aryl groups with substituent types which tend to be small and nonhydrogen bond donating (Figure 21). Lepidopteran activity is enhanced with A-ring substitution at the 4-position with 1–2 carbon lipophilic groups or, alternatively, with a 2,3- or a 2,[3,4]-ring substitution pattern. B-ring patterns are less specific, but substitution in the 2-, 2,5-, 3,5-, or 3,4,5-positions can be favorable. Coleopteran activity, on the other hand, is optimized only with the most parsimonious selection of one or

228 Ecdysteroid Agonists and Antagonists

Figure 21 Diacylhydrazine pharmacophore for two insect orders.

p9010

two small groups, exemplified by halofenzozide having a single chlorine at the A4 position. The known binding pose of the DAH chemotype in HvEcR (Billas et al., 2003) is a folded conformation with P-helicity (vida infra), wherein the t-butyl group points downward approximately toward the H6/7 loop and the A-ring is orientated towards the H7/10 interface. The A-ring ortho-methyl substituent of the studied ligand, BY-106830, is orientated to its exterior. Hydrogen-bonding interactions are made with the A-ring carbonyl (N504), B-ring carbonyl (T343), and NH group (Y408). The t-butyl group lies in a hydrophobic pocket comprising residues from H3, H11, the H6/7 loop and the H11/12 loop (Billas et al., 2003). Overall, this binding pose is quite unusual and considerably shifted in position relative to ligands of other nuclear receptors, which, when viewed together, maintain a fairly consistent center of mass more to be likened to PoA bound in HvEcR. EcR crystal structure-naive docking studies have generally placed DAHs in a more conventional position (Wurtz et al., 2000; Kumar et al., 2002). The DAHs are generally globular and do not exhibit completely dissymmetric and pronounced electrostatic surface markings. Thus, in homology model docking studies, DAH orientation tends to be multimodal. It remains to be seen what DAH docking to a genuine EcR structure will reveal. The conclusions from qualitative SAR, QSAR studies, and Lepidopteran pharmacophore analysis depicted in Figure 21 are consistent with the DAH– protein interactions implicated by the HvEcR/BY106830 crystal structure. The authors attribute the Lepidopteran specifity of the DAH BY-106830 to the lepidopteran-conserved residue, V384, which is complementary in shape, hydrophobicity and particularly the size of the 3,5-dimethyl substitution of the BY-106830 B-ring. In other insect orders, methionine occurs at this residue position and is prohibitively sterically demanding (Billas et al., 2003). It will be very interesting to see in

Figure 22 The amidoketone (AMK) generalized and lead structures.

the future just how DAHs actually do bind to nonlepidopteran EcRs. 3.4.9.3. Amidoketone SAR

The amidoketones (AMK) have been disclosed recently as an EcR-active chemotype (Tice et al., 2003a, 2003b). These compounds invoke an isosteric replacement of the B-t-butyl amide in the DAHs with a disubstituted ketone, thereby nullifying the long-held precept that nitrogens are required in the backbone. The AMKs are effective as gene switch inducers in systems based on BmEcR/RXR (B. mori) in HEK cells or on CfEcR/b-RXR-LmUSP (C. fumiferana, L. migratoria) in Chinese hamster ovary cells. AMK SARs for these two receptors indicate a pattern reflecting that found for the DAHs. Notably, the C-module, in which the Cfor-N replacement is located, shows a strong preference for four or five attached carbons, either as two acyclic susbstituents or more preferably, as a five- or six-membered ring. The most active reported representative bears a 3,5-dimethyl substitution on the B-ring, a cyclohexyl group as the C-module, and a dioxan ring fused to the A-aryl group. The crystal structure of an AMK analog of methoxyfenozide shows a remarkable fit of its backbone atoms with those of a crystal structure for GS-ETM (RG-102240, DAH: A ¼ 2-ethyl, 3-methoxy; B ¼ 3,5-dimethyl) (Figure 22) (Tice et al., 2003b). Thus, the doubly substituted backbone carbon,

p0315

Ecdysteroid Agonists and Antagonists

isosteric to N(t-Bu), appears also to serve quite adequately as a conformational determinant. 3.4.9.4. Tetrahydroquinolines

The tetrahydroquinolines (THQ) are another distinctly different ecdysonergic chemotype for which an SAR is emerging. Investigators at FMC Corp., who made the initial discovery (Chaguturu et al., 1999; Dixson et al., 2000), have reported that representatives bind competitively with PoA to the EcR, and that they have a nanomolar affinity for the Drosophila receptor, exceeding their submicromolar affinity for HvEcR, the receptor from the tobacco budworm, Heliothis virescens. FMC Corp. has also demonstrated the compounds to be insecticidal. The optimized compounds bear small alkyl or halo

229

groups in the meta- and/or para-positions of the D-ring and fluorine- or no substitution in the A- and C-rings. Subsequently at RheoGene, Smith et al. (2003) examined a designed library of THQs in an inducible gene regulation system based on AaEcR (from yellow fever mosquito, Aedes aegypti) in 3T3 cells and demonstrated that similarly optimized structures were nearly as potent as GS-ETM as gene-switch elicitors, with induction levels reaching several hundred-fold above background (Figure 23). Published data do not quite yet allow definition of a THQ pharmacophore or a QSAR. 3.4.9.5. Other Chemotypes and Nonecdysonergic Effects

Other chemotypes reported to be E agonists or antagonists such as DTBHIB, maocrystal E, or the brassinosteroids, cucurbitacins, stilbenoids, phenylalkanoids, and harpagides are not yet sufficiently represented with an adequate span of potency for satisfactory application of QSAR. In one or two cases, true ecdysonergic properties remain to be adequately demonstrated. Only a preliminary SAR examination of the electrophysiological response of taste receptors to ecdysteroids has been conducted. 3.4.9.6. Interchemotype Comparisons

Figure 23 The structure.

tetrahydroquinoline

(THQ)

generalized

The DAH chemotype is striking not only owing to its extraordinary suitability as a crop protection agent, but also owing to the utter dissimilarity of its chemical structure to the natural ecdysteroids (Table 5).

Table 5 Diacylhydrazine and ecdysteroid structures, physiochemical properties, and numbering systems

Shape Volume (A˚3) Surface area (A˚2) Polar surface area (A˚2) C log P H bond donors H bond acceptors Effective rotatable bonds Polarizability (m)

RH-75992 /tebufenozide

Ponasterone A

Globuar 365 465 47 4.51 1 2 2 7.79

Crescent 454 633 121 0.99 5 6 5 3.72

Reproduced with permission from Hormann, R.E., Dinan, L., Whiting, P., 2003. Superimposition evaluation of ecdysteroid agonist chemotypes through multidimensional QSAR. J. Comp. Aided Mol. Des. 17 (2–4), 135–153; ß Kluwer Academic Publishers.

230 Ecdysteroid Agonists and Antagonists

p0335

p0340

Accordingly, interchemotype comparisons have been undertaken in the area of physiological properties, structural superimposition and model building and, ultimately, might take place in the arena of comparative EcR crystal structures containing alternate chemotypes. In EcR competitive binding studies and tissue activation, lepidopteran-optimized DAHs such as tebufenozide and methoxyfenozide match or exceed PoA in potency, but even the prototype RH-5849 compares favorably with the less active 20E (Dhadialla et al., 1998; Nakagawa et al., 2000b). Otherwise, for other insect orders, the DAHs are generally relatively weaker. The relationship is a bit less clear in engineered inducible gene regulation systems. In the VgEcR/RXR geneswitch, which is based on the Drosophila receptor (DmEcR), nine out of 21 tested DAH ligands showed an ability to actuate this ecdysteroidresponsive switch transiently transfected into CV-1 cells, albeit with considerably less potency than muristerone A (Saez et al., 2000). On the other hand, Chinese hamster ovary cells stably transfected with the DmEcR-derived VgEcR/RXR switch and a b-galactosidase reporter responded to GS-ETM at a level comparable to PoA and muristeroneA (MuA) (Carlson et al., 2001a). In gene switches derived from lepidopteran EcRs, DAHs are generally superior to the steroids as ligands (Kumar et al., 2002). The geometrical relationship of ecdysteroids and DAHs is particularly interesting, not only because of their structural dissimilarity, but also owing to the compelling evidence, solidified by holo-EcR crystal structures, that the same binding cavity recognizes both chemotypes. This is the EcR paradox: the simultaneous high affinity for both the extended, polar, hydroxyl-laden ecdysteroids and the relatively compact, lipophilic, hydrogen bond donor-deficient diacylhydrazines. An understanding of steroid–DAH superimpositions is greatly facilitated by an appreciation for the DAH and ecdysteroid conformational space. DAHs distribute into four conformational clusters based on the E- or Z-configuration of the amide bond. Each of these is described by a trivial nomenclature (Figure 24). Furthermore, each conformational cluster exists in two enantiomorphs, depending upon the M- or P-helicity of the N–N bond (Figure 25). Since the conformation-determining amide bonds are in the backbone of the structure, each of these clusters vary quite considerably in overall shape and potential superimposition with the ecdysteroids. DAH conformational analysis by several investigators leads to the general conclusion that the extended, but especially the folded and stacked, conformations are more favorable than the hooked

Figure 24 Diacylhydrazine conformational clusters. (Reproduced with permission from Hormann, R.E., Dinan, L., Whiting, P., 2003. Superimposition evaluation of ecdysteroid agonist chemotypes through multidimensional QSAR. J. Comp. Aided Mol. Des. 17 (2–4), 135–153; ß Kluwer Academic Publishers.)

f0120

Figure 25 Diacylhydrazine enantiomorphs (folded conformation). (Reproduced with permission from Hormann, R.E., Dinan, L., Whiting, P., 2003. Superimposition evaluation of ecdysteroid agonist chemotypes through multidimensional QSAR. J. Comp. Aided Mol. Des. 17 (2–4), 135–153; ß Kluwer Academic Publishers.)

f0125

conformation. However, depending upon substitution pattern and calculation method, all four clusters can appear within as little as 4 kcal of the global minimum (Hormann et al., 2003). More accurate calculations will certainly result with application of improved N–N torsional parameterization (Reynolds and Hormann, 1996; Chakravarty et al., 1998). Perhaps owing to the fact that the folded conformation is frequently observed in crystal structures, all DAH–steroid superimpositions so far proposed, as well as all multidimensional QSAR studies, have focused on the folded conformation with either the M- or the P-helicity. The DAH conformation observed in the HvEcR crystal structure

Ecdysteroid Agonists and Antagonists

p0355

is indeed a folded conformation of P-helicity (Billas et al. 2003). The ecdysteroid conformational space varies primarily in the A-ring conformation and the trajectory of the side chain relative to the D-ring. The A-ring may take on a chair, half-chair, or twist-boat conformation; the chair conformation is found in the E crystal structure, which, in turn, is the basis for much of the modeling that has been performed. The C16/C17/C20/C22 and C17/C20/C22/C23 torsions are most conformationally-determinant; in the crystal structure, these take on values of 62.8 and 73.9 , respectively (Huber and Hoppe, 1965), thereby conferring a crescent shape to the entire molecule. Similar conformations often appear as the favored ones in 4D-QSAR studies, in which conformation is self-optimizing. Conformations incorporating a more acute bend at the core/side chain junction appear in at least one CoMFA study (Nakagawa et al., 1998) and also for some weakly active steroids in 4D-QSAR studies. Satisfyingly, the PoA conformation observed in the HvEcR crystal structure shows an A-ring in the chair conformation and an overall crescent shape. DAH/ecdysteroid superimpositions hypotheses have been derived from or analyzed by simple pharmacophore considerations (Shimizu et al., 1997), perceived substructure analogy (Mohammed-Ali et al., 1995; Sawada et al., 2003a), conformational studies (Qian, 1996), homology model docking (Wurtz et al., 2000), and multidimensional QSAR (Nakagawa et al., 1995a, 1998; Hormann et al., 2003). The latter two approaches potentially offer some degree of validation. The first superimpostion tested in a QSAR model relied on a set of 37 DAHs and six ecdysteroids tested in the previously mentioned Chilo integument assay. Overlaps were established by aligning the DAH carbonyl oxygens with O6 and O14 of the steroid, or alternatively, O14 and O20. Models were evaluated in part on the q2 values which ranged from 0.393 to 0.472. The favored model (q2 ¼ 0.472, r2 ¼ 0.892, s ¼ 0.250, five components) involved an O14/O22 alignment with the DAH B-ring oriented toward the side chain. Shimizu et al. (1997) later discovered that replacement of the DAH A-ring with simple C5–C6 alkyl chains resulted in structures with potency approaching that of tebufenozide in the C. suppressalis integument assay, and, considering the sensitivity of steroid potency to O22 configuration, hypothesized that DAH carbonyl alignments with O20 and O22 might result in superior superimpositions (Nakagawa et al., 1998). CoMFA models of this superimposition type (Figure 26) compare favorably with models based on the O14/O22 alignment

231

Figure 26 Favored superimposition for DAH–ecdysteroid CoMFA model of Nakagawa (1998). 20-Hydroxyecdysone and tebufenozide are depicted. (a) View from the b-face; (b) view towards the C6/C14 edge of the steroid. (Reproduced with permission from Nakagawa, Y., Hattori, K., Shimizu, B.-I., Akamatsu, M., Miyagawa, H., et al., 1998. Quantitative structure–activity studies of insect growth regulators XIV. Threedimensional quantitative structure–activity relationship of ecdysone agonists including dibenzoylhydrazine analogs. Pestic. Sci. 53, 267–277.)

(q2 ¼ 0.359–0.409 and 0.307–0.388, respectively), especially in view of the fact that 14OH is not necessary for receptor binding (Cherbas et al., 1982). The analyses were performed with 56 DAHs and six ecdysteroids using C. suppressalis integument data as the target variable. In another study, Drosophila BII data for 97 DAHs and 66 ecdysteroids were fitted to eight different superimpositions, representing both M and P DAH enantiomorphs, using CoMFA and 4D-QSAR in a consensus strategy (Hormann et al., 2003). The resultant QSAR models were evaluated on the basis of q2 and test set r2, using a test set comprising 19 DAHs and 33 ecdysteroids which had not been used in model construction. The best test set r2 values, generally a more rigorous criterion of model quality than training set q2, were obtained from a common superimposition (q24D-QSAR ¼ 0.722; q2CoMFA ¼ 0.607, test set r24D-QSAR ¼ 0.382, test set r2CoMFA ¼ 0.428) (Figure 27) which fortuitously resembled that obtained from a CtEcR homology model built from RAR-g (Wurtz et al., 2000). In this case, the DAH B-carbonyl aligns with the steroid B-ring carbonyl, but the DAH A-ring carbonyl does not

232 Ecdysteroid Agonists and Antagonists

Figure 27 One of the favored DAH/ECD superimpositions depicted as its 4D-QSAR model relative to PoA and tebufenozide in their postulated active conformations with respect to each model. See explanatory text for Figure 17. (Reproduced with permission from Hormann, R.E., Dinan, L., Whiting, P., 2003. Superimposition evaluation of ecdysteroid agonist chemotypes through multidimensional QSAR. J. Comp. Aided Mol. Des. 17 (2–4), 135–153; ß Kluwer Academic Publishers.)

align with any oxygen functionality. Another highly ranking superimpostion invoked a DAH carbonyl alignment with O20 of the steroid. None of these DAH/ecdysteroid superimposition hypotheses correspond to the overlap found for the HvEcR crystal structures (Billas et al., 2003). As previously alluded to, there is a structural reason to speculate that the excluding volume of M384 in H5 of DmEcR might possibly preclude this possibility altogether. Whatever the case may be, the HvEcR/BY106830 structure offers a revealing answer to the EcR paradox – the recognition of widely disparate DAH and ecdysteroid chemotypes. In the HvEcR crystal structures, this is accomplished by (1) use of only partially overlapping loci (steroid sidechain and DAH B-ring/B C›O/t-Bu), (2) use of w only one common pharmacophore element (hydrophobe corresponding to distal portion of steroid side-chain and DAH B-ring/t-Bu), and (3) interaction with common hydrogen-bonding protein residues from substantially different positions (Y408 with steroid-20-OH/DAH-NH and T343 with steroid-14-OH/DAH B C›O). It is becoming w apparent that the complete set of pharmacophore elements recognized by EcR is much more expansive than those found in the ecdysteroids alone.

One must be mindful of the possibility of multiple binding modes across EcRs from different species, across analogs of the same chemotype, or even for a single ligand with respect to a single responsive EcR. Whether one adheres strictly to a crystal structure superimposition or speculates on alternate binding modes, superimposition exercises are useful because the resulting models can be used as virtual screens with quite different characteristics than scored docking. Superimpositional studies of the alternate chemotypes are less advanced. The direct structural and SAR similarity of the AMKs strongly suggests that their steroid superimposition could be inferred from DAH considerations. The THQs, on the other hand, require an independent analysis. One illustrative possibility, generated from the genetic algorithm similarity program (GASP) algorithm, is shown in Figure 28 ( Jones et al., 1995). One might also speculate that the THQ A/D-rings or C/D-rings coincide with the DAH aryl rings. 3.4.9.7. Future Developments

QSAR methodology is important to ligand design because it enables conceptualization and succinct articulation of pharmacophoric design elements.

p0370

p0375

Ecdysteroid Agonists and Antagonists

Figure 28 A possible THQ/ECD superimposition of the fully unsubstituted 2R,4S-THQ and PoA. (a) View from the b-face of PoA; (b) view towards the C6/C14 edge of PoA.

These design elements can take the form of localized physiochemical properties, hydrogen bonding potential, shape, molecular dynamics, and whole molecule properties. These features are frequently difficult or impossible to express in simple diagrams or prose, particularly in the area of multichemotype superimposition. Applied rigorously and creatively, QSAR can evoke novel perceptions of the ligands – perceptions that are useful for further design. These principles have held true for QSAR applied to ecdysteroid agonists. Nevertheless, certain issues remain outstanding. Undersized training sets, narrow structural variation, and inadequate or imbalanced distribution across the target variable can lead to unstable models with limited explanatory power. The issue of model validation also requires more attention (Golbraikh and Tropsha, 2002). Models with low q2 values or the absence of a test set leave the question of predictability completely unanswered and often in doubt. Moreover, even where test sets are applied, the test set selection may demonstrate only that the model is interpolative rather than extrapolative. The latter is preferred. Examples of all of these compromising situations exist in the field of ecdysteroid agonist QSAR. To be sure, some of the aforementioned criteria have developed fairly recently as QSAR methodology has matured. Even applied imperfectly, the QSAR approach so far has exerted explanatory power and provided increased confidence during grueling synthesis programs. The indisputable mark of success will be a QSAR-based ligand design that would otherwise have been impossible. Multichemotype superimposition experiments may be headed in this

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direction. The demonstration that a structurally divergent active chemotype could be designed as a direct and unique consequence of a superimposition would be compelling indeed. In the future, one may anticipate more extensive use of validating test sets and a more liberal application of consensus QSAR. The CoMFA/4D-QSAR studies open this door. Parallel models using topological descriptors or alternate variable selection and statistical methods may be especially effective in combination with multidimensional QSAR; a model using topological descriptors has been built on the ecdysteroid BII data set (Golbraikh et al., 2001). Additionally, judicious application of lateral QSAR using the same training sets, but different test organisms, may shed additional light on the crossspecies selectivity question. One may also see improvements in multidimensional QSAR methodology which speak directly to the issue of multichemotype models. The availability of an EcR binding domain crystal structure, in addition to answering numerous questions in structural biology, enables de novo ligand design. It is also possible that extracted protein information can be incorporated into ligand QSAR models (Santos-Filho et al., 2001), which might be more reliably quantitative than pure docking. Both crystal structure and advanced receptor-independent QSAR models can be used as virtual screens to discover new ligands. In doing so, one must keep firmly in mind the possibility of unexpected or multiple binding modes, either within a single ligandbinding pocket or across species. It may turn out that protein dynamics simulations enhance the already illuminating perspective offered by static EcR crystal structures in explaining the level of receptor plasticity crucial for recognizing ligands of disparate structure.

3.4.10. Applications and Prospects 3.4.10.1. Insecticides

The wide diversity of chemistries that have been shown to interact with ecdysteroids receptors give hope that further commercially viable compounds can be developed as effective and selective insecticides. The lead given in this area by the DAHs is very encouraging. These compounds are chemically simple, effective, nontoxic to vertebrates, and even show some selectivity towards particular insect orders (Hsu et al., 1997; Dhadialla et al., 1998; Carlson, 2000; Carlson et al., 2001b). The discovery of the DAHs was somewhat serendipitous, but the development of further classes of ecdysteroid signaling pathway-active compounds can be facilitated

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through mechanism-based assays, QSAR, molecular modeling, and virtual screening. Prior to the identification of the DAHs, it was questioned whether ecdysteroid agonists would make effective insecticides, since it was expected that such compounds would only have an effect at the next molt. In the event, the fact that affected insects do not immediately die is not a problem because a further effect of the ecdysteroid agonist is to bring about a cessation of feeding. Ecdysteroid antagonists might be expected to be effective against pest species where the adult is the destructive stage or where reproductive disruption is sought. However, as our understanding of the pleiotropic physiological consequences of ecdysteroid (ant)agonists is still very incomplete, we should not exclude the possiblitiy that ecdysteroid antagonists may also be effective against larval insects or embryos. 3.4.10.2. Genetic Modification of Ecdysteroid Levels in Crop Plants

Very few crop species accumulate ecdysteroids. Only spinach (Spinacia oleracea; Chenopodiaceae) and quinoa (Chenopodium quinoa; Chenopodiaceae) accumulate significant amounts (Dinan, 1992, 1995b). However, available evidence indicates that most, if not all, higher plant species possess the genetic capacity to produce ecdysteroids, but that the pathway is downregulated (Dinan et al., 2001d). If this is true, this will considerably facilitate the elevation of ecdysteroid levels in crop species, as alteration of the regulation will result in changes in ecdysteroid level. This could require alteration of only one gene, rather than the incorporation of a series of genes for the entire biosynthetic pathway. As an absolute prerequisite to the genetic modification of ecdysteroid levels in crop species, it is necessary that the biosynthetic pathway for phytoecdysteroids should be elucidated. Significant progress is now being made in this area using hairy root cultures of A. reptans (Fujimoto et al., 2000), which produce ecdysteroids in large amounts and allow stable isotope-labeled precursors to be incorporated in high yield permitting analysis of ecdysteroidal products by 13C NMR. The generality of the Ajuga pathway(s) will need to be assessed and the regulation of the pathways determined in order to identify the step(s) which are downregulated in nonaccumulating species. Ecdysteroid-accumulating lines of crop species could be generated by genetic engineering approaches or by traditional plant breeding techniques. Ultimately, it should be possible to modify the profile of ecdysteroids present in crop plants to maximise resistance against even polyphagous pest species which possess particular detoxification

mechanisms against common major ecdysteroids (e.g., 22-acylation of 20E) and/or to elevate levels of ecdysteroids of higher biological activity (e.g., PoA). The very low vertebrate toxicity (and even benefit as nutritional supplements) and lack of taste of ecdysteroids would mean that phytoecdysteroid accumulation need not be restricted to the portions of the plant not to be consumed. 3.4.10.3. Gene Switching Elicitors

Engineered inducible gene expression systems are gaining research attention and technological significance. Gene switches permit the temporal and/or spatial expression of transfected genes, with potential application in medicine (gene therapy, tissue engineering, biotherapeutic protein production), agriculture (crop protection, yield enhancement, nutritional enhancement, pharmaceutical production) and in basic research for investigation of gene function (proteomics and drug lead discovery). Several different approaches are being developed (Fussenegger, 2001) and the use of native or hybrid ecdysteroid receptors with ecdysonergic ligands is one of the current forerunners in the field. The elicitor should be effective at low concentrations and should be target-specific (i.e., nontoxic). For medical applications, the elicitor should be nonallergenic and preferably orally available. The elicitor can also affect dynamic properties of the gene switch such as response time, cycling, dynamic range, maximum response, span of the dose-response curve, and tissue specificity. These properties should be tailored to the specific application. Ecdysteroids and DAHs, which, taken together, span a wide range of physiochemical properties, and are both highly promising chemotype candidates for many gene-switch applications. The gene-switch potency, absorption, distribution, and known pharmacological properties of the ecdysteroids are favorable toward most uses of a gene switch. On the other hand, metabolism and elimination may be mitigating factors. When native or hybrid ecdysteroid receptors are transfected into mammalian, plant or yeast cells, the EcRs possess altered ecdysteroid specificity and potency. In some cases, 20E is hardly active at all and PoA and MuA are required at concentrations 100-fold higher than in insect systems. More extensive SAR or even QSAR studies could assist identification of the most potent ecdysteroids for these systems. A detailed discussion of ecdysteroid-regulated gene switches is beyond the scope of this review, but has been reviewed elsewhere recently (Lafont and Dinan, 2003). The properties of DAHs that are quite favorable to gene switch applications are potency towards

Ecdysteroid Agonists and Antagonists

lepidopteran EcRs, metabolic stability, chemical simplicity, and absence of toxicity. Available data suggest that absorption and distribution properties may be acceptable for pharmaceutical applications. However, solubility is potentially a barrier to be overcome. It remains to be seen if the DAH physical properties are an issue for agricultural gene switch applications in which plant uptake requirements may differ from those needed for the already successful insecticidal applications. An alternative DAH (GS-ETM) (Figure 6) has been developed for use with mammalian cells. QSAR has been brought to bear on DAH potency in a mammalian cellular EcR-based gene switch system (Hormann et al., 2002), and should aid future studies that pertain to pharmacokinetics. Orthogonal gene switches, that is to say, multiple parallel switches which can be actuated simultaneously, noninteractively, and independently, would constitute a significant enhancement to the field of engineered gene regulation. A major potential benefit of EcR systems is their inherent orthogonality to the corresponding transcriptional machinery of the mammalian host. A concomitant liability, however, is the potential for immunogenicity. Likewise, ecdysteroids are not regarded as normal, interactive components of mammalian cells and may have a natural built-in orthogonality. However, the ecdysteroids are present in certain plants as phytoecdysteroids, can be present in the human diet, and may later turn out to have unacceptable pleiotropoic effects. For purposes of orthogonality, the DAHs look attractive. Certainly, the order-specificity of DAHs can be exploited to develop switches for different hybrid ecdysteroid receptors in the same system. For representative ecdysteroids, an otherwise ecdysteroid-responsive EcR has been rendered ecdysteroid-refractive by sitedirected mutagenesis, auguring well for orthogonal systems which also utilize a DAH elicitor in a parallel channel, as has been addressed at RHeoGene, L.L.C. (Kumar et al., 2002). 3.4.10.4. Human Health Preparations and Animal Food Supplements p0435

The effects of ecdysteroids on mammals have recently been reviewed (Lafont and Dinan, 2003), to which the reader is referred for more extensive treatment of this topic. As a consequence of the reputed anabolic effects of ecdysteroids on animals, a large number of ecdysteroid-containing preparations are now available for use by sportsmen and body-builders. While there is evidence for the positive anabolic effects of ecdysteroids on Japanese quail (Koudela et al., 1995; Sla´ ma et al., 1996) and

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pigs (Selepcova et al., 1993; Kratky et al., 1997), the effects are not dramatic (ca. 15% increase in body mass over controls). Ecdysteroids may also improve physical performance, as demonstrated on rats (Chermnykh et al., 1988). Evidence has been provided for effects of ecdysteroids on cellular proliferation, protein synthesis, glucose metabolism, lipid metabolism, nerve function, hepatic function, cardiovascular function, renal function, and the immune system, indicating that ecdysteroids might contribute to the relief of conditions such as diabetes, hypercholesterolemia, wound healing, mental disorders, myocardial arrhythmia, inflammations, etc. (review: Lafont and Dinan, 2003). All these aspects need further study and verification, but the lack of toxic dose required to kill 50% of the organism (LD50 > 6 g kg
1), androgenic or (anti)estrogenic effects make pharmacological applications of ecdysteroids feasible and potentially attractive. 3.4.10.5. Improvement of Silk Yields

Exogenous ecdysteroids fed to larvae of the silkworm Bombyx mori at certain stages of development enhance synchronous development of the larvae and, when coadministered with a juvenile hormone analog, significantly elevate the yield of silk obtainable from the cocoon (Chou and Lu, 1980; Ninagi and Maruyama, 1996). Such treatments have considerable potential to improve the efficiency of silkworm-rearing by Asian farmers, especially since the hormonal treatments could be based on extracts of appropriate indigenous plant species (Chandrakala et al., 1998). 3.4.10.6. Endocrine Disruption

Although it is recognized that endocrine disruption in invertebrates may be a potentially serious problem, many of the necessary experimental tools are not available for research in this area. The many gaps that still exist in our knowledge of invertebrate endocrinology limit the ability to interpret the toxic effects of exogenous chemicals. Even for the insects, where quite extensive endocrinological information is available for a number of model species, they do not correspond to aquatic species or to the sentinel species one would wish to use to monitor for EDCs. Thus, it is to be expected that there will be a revival in interest in hormone biochemistry for environmentally relevant species to provide baseline information against which exposed insects can be compared. Further, with the sequencing of the D. melanogaster genome and the development of DNA arrays, it is to be expected that the influence of exogenous chemicals on gene expression will be investigated more extensively.

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3.4.10.7. QSAR and Molecular Modeling

If we are to understand the role of phytoecdysteroids in deterring insect predation, the QSAR/modeling approach should be extended to more analogs and to more species and to the SAR of the deterrent antifeedant/toxic effects of ecdysteroids. This would go some way to explain the diversity of phytoecdysteroid structures in plants. The pharmacological effects of ecdysteroids (almost exclusively 20E) on mammals are only now being accepted and it seems that there are many potential beneficial effects. In the future, it will be helpful to conduct QSAR and docking/scoring studies to identify the most effective analogs. In the area of gene switching using ecdysteroids as analogs, SAR will be important not only because hybrid ecdysteroid receptors in mammalian cells have altered specificities, but also because ligands will need to be tailored as specific elicitors for multiplex systems expressing several receptor types simultaneously, in order to enable selective activation of their regulated genes.

Acknowledgments We are grateful to Y. Nakagawa for molecular structures of diacylhydrazine–ecdysteroid superimpositions. Research conducted at Exeter University was supported by the BBSRC, Rohm & Haas Co., RheoGene Inc., Astra-Zeneca plc, EU-INTAS and the Leverhulme Trust.

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a-acylaminoketone ecdysone agonists for control of gene expression. Bioorg. Med. Chem. Lett. 13, 1883–1886. Tomaschko, K.-H., 1999. Nongenomic effects of ecdysteroids. Arch. Insect Biochem. Physiol. 41, 89–98. Toya, T., Yamaguchi, K., Endo, Y., 2002. Cyclic dibenzoylhydrazines reproducing the conformation of ecdysone agonists, RH-5849. Bioorg. Med. Chem. Lett. 10, 953–961. Velgova´ , H., La´bler, L., Cˇ erny´, V., Sˇ orm, F., Sla´ ma, K., 1968. On steroids. CXI. Some further compounds producing molting deficiencies in an insect. Coll. Czech. Chem. Commun. 33, 242–256. Voigt, B., Whiting, P., Dinan, L., 2001. The ecdysteroid agonist/antagonist and brassinosteroid-like activities of synthetic brassinosteroid/ecdysteroid hybrid molecules. Cell. Mol. Life Sci. 58, 1133–1140. Wang, S.F., Ayer, S., Segraves, W.A., Williams, D.R., Raikhel, A.S., 2000. Molecular determinants of differential ligand sensitivities of insect ecdysteroid receptors. Mol. Cell. Biol. 20, 3870–3879. Wing, K.D., 1988. RH 5849, a nonsteroidal ecdysone agonist: effects on a Drosophila cell line. Science 241, 467–469. Wing, K.D., Slawecki, R.A., Carlson, G.R., 1988. RH5849, a nonsteroidal ecdysone agonist: effects on larval Lepidoptera. Science 241, 470–472. Wurtz, J.-M., Guillot, B., Fagart, J., Moras, D., Tietjen, K., et al., 2000. A new model for 20-hydroxyecdysone and dibenzoylhydrazine binding: a homology modeling and docking approach. Protein Sci. 9, 1073–1084. Xu, Y., Fang, F., Chu, Y., Jones, D., Jones, G., 2002. Activation of transcription through the ligand-binding pocket of the orphan nuclear receptor ultraspiracle. Eur. J. Biochem. 269, 6026–6036. Yao, T.P., Forman, B.M., Jiang, Z.Y., Cherbas, L., Chen, J.D., et al., 1993. Functional ecdysone receptor is the product of EcR and Ultraspiracle genes. Nature 366, 476–479.

3.5 The Ecdysteroid Receptor V C Henrich, University of North Carolina-Greensboro, Greensboro, NC, USA ß 2005, Elsevier BV. All Rights Reserved.

3.5.1. Overview 3.5.1.1. Molecular Identity of the Insect Ecdysteroid Receptor 3.5.1.2. Historical Perspective 3.5.1.3. Chapter Organization 3.5.2. Ecdysteroid Receptor Structure, Biophysics, and Biochemistry 3.5.2.1. Domain Organization and Amino Acid Alignment 3.5.2.2. Strategies for Identifying Insect Receptors 3.5.2.3. Homology Models and Crystal Structure for Ecdysteroid Receptor Components 3.5.2.4. Biochemical Analysis of EcR and USP 3.5.3. Functional Characterization of the Ecdysteroid Receptor 3.5.3.1. Cell Culture Studies: Rationale 3.5.3.2. Cell Cultures: Characterization 3.5.3.3. Cell Cultures: Functional Receptor Studies 3.5.3.4. Ecdysteroid-Inducible Cell Systems 3.5.4. Cellular, Developmental, and Genetic Analysis 3.5.4.1. The Salivary Gland Hierarchy: A Model for Steroid Hormone Action 3.5.4.2. Molecular and Genetic Characterization of the Puff Hierarchy 3.5.4.3. Developmental Regulation of Ecdysteroid Response in D. melanogaster 3.5.4.4. Conservation of Ecdysteroid Response Among Insects 3.5.4.5. In Vivo Molecular and Genetic Analysis of EcR and USP 3.5.4.6. Ecdysteroid Receptor Cofactors 3.5.4.7. Orphan Receptor Interactions with EcR and USP 3.5.4.8. Ecdysteroid Action and Other Developmental Processes 3.5.4.9. Juvenile Hormone Effects upon Ecdysteroid Action 3.5.5. Prognosis

3.5.1. Overview 3.5.1.1. Molecular Identity of the Insect Ecdysteroid Receptor

The morphogenetic events associated with insect development are largely triggered by the action of a single class of steroid hormones, the ecdysteroids (see Chapter 3.3).1 Among all insect orders examined so far, it has been established that the ecdysteroid-induced orchestration of molting and metamorphosis is mediated by a heterodimer comprised of the ecdysone receptor (EcR; Koelle et al., 1991) and Ultraspiracle (USP; Oro et al., 1990; Shea 1 Ecdysteroids refers to the family of ecdysteroids including natural and artificial steroids. Individual forms, including the classic active ‘‘molting hormone,’’ 20-hydroxyecdysone (20E, formerly b-ecdysone), will be specified throughout the chapter, as will its precursor, ecdysone (formerly a-ecdysone). Ecdysteroid agonists refers to molecules which are not steroidal, but which are capable of inducing one or more insect EcR genes in a given experimental regime.

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et al., 1990; Henrich et al., 1990) that is stabilized by 20-hydroxyecdysone (20E) and which recognizes specific promoter elements in the insect genome to regulate transcription (Yao et al., 1992, 1993; Thomas et al., 1993). Both proteins belong to a superfamily of nuclear receptors which mediate transcriptional responses to steroids and other lipophilic molecules (Blumberg and Evans, 1998) (see Chapter 3.6). Recently, a second ecdysteroidsignaling pathway involving USP and another nuclear receptor DHR38 (Sutherland et al., 1995) has been discovered (Baker et al., 2003). The insect EcR is a distant relative of the vertebrate farnesol X receptor (FXR; Forman et al., 1995) and LXR (Willy et al., 1995). EcR was first identified in the fruit fly, Drosophila melanogaster, and has since been found in several insects, in the ixodid tick (Amblyomma americanum; Guo et al., 1997) and in the fiddler crab (Uca pugilator; Durica et al., 2002). No functional EcR ortholog has been reported outside the arthropod phylum (Bonneton

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et al., 2003) (see Chapter 3.6). The Ultraspiracle (USP) protein is an ortholog of the retinoid X receptor (RXR), which in turn is a heterodimeric partner for several other nuclear receptors (Mangelsdorf et al., 1990; Oro et al., 1990). Among noninsect invertebrates, an RXR ortholog has been reported from A. americanum (Guo et al., 1998), U. pugilator (Chung et al., 1998), and the parasitic nematode, Dirofilaria immitis (Shea et al., 2004). RXR is widely found in humans and other vertebrates, reflecting its early evolutionary origins (see Chapter 3.6). Both EcR and USP have diverged evolutionarily among the insect orders (see Chapter 3.6), indicating that the functional properties of EcR and USP have also diverged among them. 3.5.1.2. Historical Perspective

The pursuit of the insect ecdysteroid receptor and its activities has led to a convergence of several basic and applied research disciplines. The ability of a single steroid hormone to induce widespread and coordinated changes in gene transcription has attracted basic scientists interested in the mechanistic basis of its variable gene and cellular action. For insect biologists, the diversity of developmental responses triggered by ecdysteroids among the insect orders reveals the essential importance of this process for understanding the evolutionary diversity and adaptability seen among individual insect species. In turn, the variety of responses among the insect orders has led to attempts to disrupt ecdysteroid receptor action with species-specific agonists and antagonists such as the bisacylhydrazines (Wing, 1988) for insecticidal purposes (Dhadialla et al., 1998, and references therein; see Chapter 3.4). Finally, because ecdysteroids exert no harmful effects in humans and other organisms (see Chapter 3.4), the ecdysteroid receptor is now viewed as a potential inducer of beneficial transcriptional responses in plant and animal cells. Much of the work on ecdysteroid action that was reported prior to 1985 appeared in the previous edition of this series and will not be repeated here (Riddiford, 1985; O’Connor, 1985; Yund and Osterbur, 1985), except for those early events which have had an obvious bearing on continuing investigations. The effects of ecdysteroids upon gene transcriptional activity became apparent through studies of the effects of ‘‘molting hormone,’’ 20E, on the polytene chromosomes of insects such as D. melanogaster and Chironomus tentans (midge). Puffing was first reported in 1933 in D. melanogaster by E. Heitz, and in the early 1950s Wolfgang Beermann performed seminal work on puff changes in C. tentans. Later, Ulrich Clever demonstrated that the

insect ‘‘molting hormone’’ induced puffing changes and thus reported the first evidence that steroid hormones act directly upon the transcriptional activity of specific target genes, now regarded as a central feature of steroid hormone action in all animals. A variety of biochemical studies also established the presence of a protein in cellular extracts that binds to ecdysteroids, further indicating that an intracellular receptor mediates the transcriptional response (O’Connor, 1985; Yund and Osterbur, 1985, and references therein). Conceptually, the polytene chromosomes can be viewed as an in situ expression microarray (Figure 1), since the puff sites disclose the transcriptional activity of specific genes (by chromosomal location rather than by sequence). This natural array has the added and unique benefit of showing temporal changes in puff size that roughly indicate continuous changes in transcriptional rate. Hans Becker noted in 1959 that ‘‘early’’ puffs appear when incubated with the ecdysteroidogenic ring gland and regress as other ‘‘late puffs’’ appear (Becker, 1959). Ulrich Clever and Michael Ashburner later showed that blocking the protein translation of early puff RNAs with cycloheximide treatment prevents the regression of some early puffs, and simultaneously prevents the appearance of many late puffs (Asburner et al., 1974). From these findings, Ashburner postulated that an intracellular

Figure 1 Prepupal chromosome III from Chironomus tentans labeled with antibodies against EcR (green) and RNA polymerase II (red). Yellow signals indicate colocalization of the two antibodies. Green signal is a fixation artifact. See Wegmann et al. (1995) for methods. (Photograph courtesy of Markus Lezzi.)

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ecdysteroid receptor directs a transcriptional response at early puff sites in the presence of 20E. Further, the early puff gene products regulate the appearance of later puffs and also feedback to repress their own expression. This original model for ecdysteroid action has proven remarkably durable over the years, though it is often overlooked that several puffs described by Ashburner have not been placed within the regulatory hierarchy and that some features of the model have not yet been pursued at the molecular level. The role of a receptor was also implicated by the identification of an ecdysone response element (EcRE) in the ecdysteroid-inducible promoter of the 27 kDa heat shock protein (hsp27) of D. melanogaster (Riddihough and Pelham, 1987). The palindromic inverted nucleotide repeat sequence of this ecdysteroid receptor target resembles the motifs that at the time of its discovery were associated with a growing class of nuclear receptors for several vertebrate hormones, including the glucocorticoids and estrogen (Hollenberg et al., 1985; Green et al., 1986), raising anticipation that the ecdysteroid receptor is evolutionarily conserved, just as the process of steroid action on transcriptional activity is mechanistically conserved. Much of the progress made on the molecular genetic basis of ecdysteroid action can be traced to 1968 when, during a sabbatical at CalTech, David Hogness met Ashburner, who was a postdoctoral associate there. Hogness later visited the laboratory of Beermann at Tubingen, and decided to identify the ‘‘puff’’ genes targeted by ecdysteroid action. The conviction to this goal was strong enough that Hogness isolated overlapping genomic DNA segments over a span of several hundred kilobases to find them. His laboratory successfully ‘‘walked’’ through the chromosomal region associated with two early puff genes using this positional cloning approach (Burtis et al., 1990; Segraves and Hogness, 1990). One of the early puff genes, E75,2 encodes a member of the nuclear hormone receptor superfamily (Segraves and Hogness, 1990), as defined by two cysteine–cysteine zinc fingers that are responsible for the receptor protein’s recognition of specific promoter elements. 2

The puff sites in Drosophila melanogaster were originally identified by their cytological location among the 102 subintervals arbitrarily delineated across the sex chromosome (X or chromosome 1, subintervals 1–20) and three autosomes (chromosome 2, subintervals 21–60; chromosome 3, subintervals 61–100, chromosome 4, subintervals 101–102) that comprise the D. melanogaster genome. The puffs are further designated as E (early or early-late) or L (late) based on their temporal appearance after 20E incubation.

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E75, several other ecdysteroid-responsive gene products in Drosophila, and numerous other superfamily members are ‘‘orphan receptors,’’ that is, they carry the zinc finger configuration but do not mediate transcriptional activity through any known ligand. When E75 was used as a probe to screen a cDNA library, one of the recovered candidate clones proved to be the D. melanogaster ecdysone receptor gene (DmEcR; Koelle et al., 1991), and a functional receptor was also recovered by biochemical purification from a Drosophila embryonic cell line (Luo et al., 1991). In reality, EcR dimerizes with a second nuclear receptor, Ultraspiracle (DmUSP; Yao et al., 1992, 1993; Thomas et al., 1993), whose structural resemblance to the vertebrate RXR suggests its conservation as a heterodimeric partner for EcR. The recovery of EcR, USP, and orphans such as E75 inspired a more exhaustive search for nuclear receptors in D. melanogaster, and the annotated genome now lists 21 nuclear receptors. Among them, only EcR unambiguously interacts physically with an activating ligand, though there is evidence that USP binds to juvenile hormone (Jones and Sharp, 1997; Jones et al., 2001). The recovery and characterization of EcR, USP, other orphan receptors, and various ecdysteroidinducible targets in other insects has proceeded at a quickening pace in recent years, and, along with it, a proliferation of DNA probes and antibodies have been developed to examine these players. Several insect researchers have actively identified orthologs for EcR and USP and its puff gene targets so that a rapidly expanding body of comparative information is appearing in the literature. The abundance of information accumulated in the Drosophila system has heavily influenced mechanistic interpretations of ecdysteroid action, though it is apparent from the comparative studies done so far that ecdysteroid action at the developmental level varies among insect species, consistent with their wide range of adaptations to the environment. Complete EcR and USP sequences have so far been reported for a few dozen species among four of the insect orders (Diptera, Lepidoptera, Orthoptera, and Coleoptera) and two other arthropod classes. If the variety of results obtained from these studies provide any indication, then only a fraction of the potential information concerning the ecdysteroid receptor has so far been retrieved and assimilated. 3.5.1.3. Chapter Organization

The straightforward thesis of the ecdysteroid hierarchy based on the chromosomal puff response seems to contrast with the complexity found in many

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in vivo experiments. It is apparent that ecdysteroid responsiveness varies widely by tissue or cell type, species, incubation conditions, developmental time, activating ligand, ligand concentration, and treatment duration, presumably because the functional capacity of the receptor itself varies in these regimes. Nevertheless, several common and conserved themes have emerged from this work that are valuable not only for insect biologists, but also for other geneticists, cell biologists, and endocrinologists. The organization of this chapter is based largely on the criteria by which the EcR of D. melanogaster was originally defined: (1) the deduced EcR amino acid sequence includes the zinc fingers and ligandbinding domain helices that typify nuclear receptors, (2) extracts of cells expressing EcR bind to 125 I-iodoponasterone and this binding disappears with the addition of anti-EcR antibodies, (3) insect cellular extracts carry a protein that binds specifically to an hsp27 EcRE and this binding disappears in the presence of anti-EcR antibodies, (4) ecdysteroid-inducible transcription is restored to an ecdysone-insensitive Drosophila cell line by transfecting the cells with EcR cDNA, and (5) the pattern of spatial and temporal expression of EcR during premetamorphic development is consistent with its role as a mediator of the ecdysteroid response (Koelle et al., 1991). This chapter will examine the information gathered about the ecdysteroid receptor at three levels, and provide brief descriptions and references for the tools used to undertake those experiments. First, the receptor will be viewed as a structural entity, with particular emphasis on the sequence characteristics of EcR and USP as well as their biophysical and biochemical properties. These studies build upon the first three properties noted for EcR in its original characterization. Second, and as in the original report, the transcriptional function of the ecdysteroid receptor will be examined, with particular regard to its interactions with ligand, other protein factors, and promoter elements. This section will also describe a few of the ecdysteroid-inducible systems which have been developed for various applications. Third, the ecdysteroid receptor will be viewed from a cellular and developmental perspective in vivo, with special emphasis on the spatial and temporal diversity of ecdysteroid-mediated action, along with the approaches used to address these questions. Finally, the chapter will offer a prognosis concerning important and unresolved problems surrounding ecdysteroid action via its receptors, along with possible experimental technologies and strategies that might clarify them.

3.5.2. Ecdysteroid Receptor Structure, Biophysics, and Biochemistry 3.5.2.1. Domain Organization and Amino Acid Alignment

Both EcR and USP belong to the superfamily of nuclear receptors first described for several steroid hormones and vitamins among the vertebrates. Like their evolutionary counterparts, EcR and USP are structurally modular, that is, they are composed of distinct domains responsible for specific molecular functions (Figure 2). Individual domains are at least partially autonomous in their function, since domains of different nuclear receptors can be swapped to create structural and functional chimeras. Chimeras derived from the EcR and USP of different insect species have been used to compare and differentiate their functional capabilities (e.g., Suhr et al., 1998; Wang et al., 2000; Henrich et al., 2000). The basic organization of nuclear receptors has been described extensively in other reviews, and will be examined here exclusively in terms of the functional features found in insects and associated with each of the domains: (1) the N-terminal (A/B) domain through which nuclear receptors interact with other transcriptional factors, (2) the DNA binding domain (DBD) or C domain which is responsible for the receptor’s recognition of specific DNA response elements in the genome and which is comprised of two cysteine–cysteine zinc fingers that define proteins as members of the nuclear receptor superfamily, (3) the hinge region (D domain), which has been implicated in ligand-dependent heterodimerization along with the E domain, nuclear localization, and DNA recognition, (4) the E- or ligand-binding domain (LBD), which is usually comprised of 12 alpha-helices that form a ligandbinding pocket, and (5) the F domain, which is found as a nonconserved sequence in all of the insect EcRs but not in any known USP or RXR sequence. 3.5.2.1.1. The A/B domain This domain is sometimes referred to as the trans-activation domain because this portion of the nuclear receptor interacts with the cell’s transcriptional machinery and is responsible for a ligand-independent transcriptional activation function (AF1). The A/B domain tends to be variable among nuclear receptors, though there is modest similarity in this portion of EcR and USP among insect species. In several species, multiple isoforms of EcR and/or USP with different A/B domains arise through the activity of alternative promoters and/or alternative pre-mRNA splicing. The occurrence of multiple isoforms for EcR and

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Figure 2 Domain organization of nuclear receptor superfamily members. Bars denote regions of the receptor associated with specific subfunctions or suspected subfunctions (as indicated with a ?). Boxed regions indicate consensus amino acid sequences for the DNA-binding domain of insect EcR and USP based on Bonneton et al. (2003). Asterisks (*) indicate positions of two sets of four cysteines which ligand to zinc ion to form each of the two zinc fingers.

USP does not follow any apparent pattern and therefore does not seem to be tied to an obvious evolutionary mechanism. For instance, so far only the DmEcR gene among insects encodes three isoforms (A, B1, and B2; Talbot et al., 1993). Among cyclorapphous flies the Mediterranean fruit fly (Ceratitis capitata, Cc) EcR gene specifies both the A and B1 isoforms (Verras et al., 2002), whereas only the B1 isoform has been found so far in the sheep blowfly (Lucilia cuprina, Lc) genome (Hannan and Hill, 1997). Among other Diptera, the mosquito (Aedes aegypti, Aa) EcR encodes the A and B1 isoforms (Wang et al., 2002), but the midge (Chironomus tentans,Ct) encodes only the B1-like isoform (Imhof et al., 1993). The same variable pattern is found among the lepidopteran species. The corn earworm (Heliothis virescens, Hv; Martinez et al., 1999a) has only EcR-B1, whereas the spruce budworm (Choristoneura fumiferana, Cf; Perrera et al., 1999a), the rice stem borer (Chilo suppressalis, Cs), the tobacco hornworm (Manduca sexta, Ms; Jindra et al., 1996), and the silkmoth (Bombyx mori, Bm; Swevers et al., 1995; Kamimura et al., 1997) express both an A and B1 isoform. An A and B1 isoform of EcR has also been recovered from the mealworm (Tenebrio molitor, Tm) genome (Mouillet et al., 1997) and a single EcR isoform has been recovered from the migratory

locust (Locusta migratoria, Lm; Saleh et al., 1998). The most complex EcR gene organization is found in a noninsect species, A. americanum, wherein at least three A/B isoforms have been identified, each of which resembles the D. melanogaster A isoform sequence (Guo et al., 1997). The functional properties of the insect EcR isoforms and their distinct developmental roles will be discussed later in the chapter, though it is noteworthy that the B2 isoform uniquely associated with D. melanogaster is also the most efficient for rescuing larval development in EcR homozygous mutants that would otherwise die during embryonic development (Li and Bender, 2000). This observation simultaneously highlights the possible uniqueness of the fruit fly EcR and the potential significance of the A/B domain for comparison among the insects. In one species, C. tentans, multiple USP isoforms differing in their A/B domain occur along with only a single EcR isoform (Vogtli et al., 1999). The migratory locust (Locusta migratoria; Lm) also expresses multiple USP isoforms along with the single EcRB1 isoform found so far, though the USP variants do not arise from differential splicing in the A/B domain (Hayward et al., 1999, 2003). Along with two EcR isoforms, two USP isoforms exist in A. aegypti (Kapitskaya et al., 1996), M. sexta (Jindra et al., 1997), and B. mori (Tzertzinis

248 The Ecdysteroid Receptor

et al., 1994). Two RXR/USP isoforms also exist in A. americanum, along with its three EcR isoforms (Guo et al., 1998). Other complete USP sequences have been reported for L. cuprina (Hannan and Hill, 2001) and C. fumiferana (Perera et al., 1998). 3.5.2.1.2. The C domain or DNA-binding domain (DBD) The members of the nuclear receptor superfamily are defined by a 66–68 amino acid region known as the cysteine–cysteine zinc finger (Figures 2 and 3). The EcR DBD closely resembles the vertebrate FXR, and both EcR/USP and FXR/ RXR recognize a palindromic inverted repeat sequence separated by a single nucleotide (IR1); the hsp27 EcRE follows this arrangement. A consensus sequence based on the investigation of several functional IR1 elements is: 50 -PuG(G/T)T(C/G) A(N)TG(C/A(C/A(C/t)Py (Antoniewski et al., 1993). As will be noted later, the EcR/USP complex is also capable of recognizing direct repeat elements

(Antoniewski et al., 1996; Vogtli et al., 1998). The DBD amino acid sequence is highly conserved among the Diptera and Lepidoptera, but specific residues are substituted in the EcR DBD of other insects (Henrich and Brown, 1995; Bonneton et al., 2003) (see Chapter 3.6). 3.5.2.1.3. The D domain or hinge region The D domain includes the amino acids which lie between the zinc fingers and the ligand binding domain (LBD). Portions of this region are highly conserved among all the insect EcRs, including a T-box motif that plays a role in DNA recognition (Devarakonda et al., 2003; Figure 3) and an A-box, a helical structure that is essential for high affinity recognition of a DNA response element (NiedzielaMajka et al., 2000). The D region of EcR is also essential for ligand dependent heterodimerization with USP (Suhr et al., 1998; Perera et al., 1999b). The USP hinge region includes a highly conserved

Figure 3 The protein constructs used for a crystallization study, designating amino acid contact sites with noted DNA interactions. (a) Sequence of Drosophila melanogaster EcR DBD with liganded zinc fingers, T-box, and A box. (b) Sequence of D. melanogaster USP DBD with liganded zinc fingers and T-box. Lower case letters designate artifacts of cloning in the constructs used for the study. (Reproduced with permission from Devarakonda, S., Harp, J.M., Kim, Y., Ozyhar, A., Rastinejad, F., 2003. Structure of the heterodimeric ecdysone receptor DNA-binding complex. EMBO J. 22, 5827–5840.)

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T-box motif, which plays a role in site recognition of DNA response elements but is not essential for heterodimerization (Figure 3; Niedziela-Majka et al., 2000; Devarakonda et al., 2003). 3.5.2.1.4. The E domain or ligand binding domain (LBD) The domain of the nuclear receptor which is responsible for interacting with the receptor’s cognate ligand is the E domain or LBD. The EcR and USP LBD fall within the canonical organization observed for almost all other members of the nuclear receptor superfamily. The region is comprised of 12 a-helices that form a ligand-binding pocket which holds the cognate ligand. The sequence of the EcR LBD is highly conserved among the insects, consistent with the widespread occurrence of the molting hormone, 20E, among the insect orders (see Figure 4). Nevertheless, the EcR LBD sequences from some insect orders have not been reported, and possibly important divergences among them remain unknown. For EcR, a liganddependent transcriptional activation function (AF2) is localized in the most carboxy-terminal helix 12, which folds over the pocket to hold the ligand molecule inside. This folding thereby creates an interactive surface with other proteins that ultimately modulates the receptor’s transcriptional activity. For nuclear receptors generally, a dimerization interface lies along helices 9 and 10 (Perlmann et al., 1996) and ligand-independent transcriptional functions (AF1) have also been associated with various portions of the EcR LBD (Hu et al., 2003). Later portions of the chapter will explore not only the functional features of the LBD based on biochemical, cellular, and in vivo experiments, but will also discuss the effects of LBD modification and speciesbased sequence differences upon EcR and USP capabilities. As noted elsewhere (see Chapter 3.6), the USP LBD has diverged considerably over evolutionary time. Whereas the USP LBD of some insect orders bears a fairly close resemblance to the vertebrate RXR, the dipteran and lepidopteran USP LBDs include a loop between helices 1 and 3 (Figure 5; Billas et al., 2001; Clayton et al., 2001). The diversity is further elaborated in the fly sequences, which include several additional, glycine-rich stretches that are not found in other insects. The aforementioned Lm USP includes short and long variants in the LBD which arise by an unidentified molecular process (Hayward et al., 2003). Among the 21 nuclear receptors in D. melanogaster, all but two of these proteins (Knirps, Nauber et al., 1988; and Knirps-related, Oro et al., 1988) carry the 12 alpha-helices that typify the LBD.

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Except for EcR and possibly USP, none of these 19 superfamily members are known to interact physically with a cognate ligand. Several show patterns of expression that suggest a role in mediating ecdysteroid-inducible responses (Sullivan and Thummel, 2003) and one, DHR38, is activated by ecdysteroids via a novel mechanism (Baker et al., 2003). 3.5.2.1.5. F domain Only a few members of the nuclear receptor superfamily possess the F domain, but the EcR family members carry up to 226 amino acids (in D. melanogaster) beyond the helix 12 region, though these are not conserved among the insect EcR sequences. Apparently, the F-domain of DmEcR is totally dispensible, since its cleavage produces a receptor that has the same transcriptional capabilities as the full-length EcR (Hu et al., 2003). Recently, a variant of the Aedes albopictus EcR has been discovered which lacks seven amino acids in this carboxy-terminal region. While the functional significance, if any, of this deviation is not known, only one of the forms normally predominates in a mosquito cell line (Jayachandran and Fallon, 2001). 3.5.2.2. Strategies for Identifying Insect Receptors

The identity of EcR described earlier by using a DNA probe derived from the E75 ‘‘early puff’’ gene quickly led to similar strategies for identifying EcR orthologs in other insects. These attempts have sometimes employed conventional probes, typically derived from the DBD sequence of DmEcR, to screen cDNA libraries (e.g., Mouillet et al., 1997; Guo et al., 1997). In many instances a combination of approaches utilizing cDNA library screening along with polymerase chain reaction (PCR) using primers from conserved EcR regions have been utilized to obtain full-length EcR cDNA clones (e.g., Swevers et al., 1995). Similarly, the use of PCR to amplify a small portion of the EcR DBD sequence, followed by cDNA library searches has also resulted in the successful recovery of EcR cDNA clones (e.g., Hannan and Hill, 1997). Because many insects encode multiple EcR isoforms, 50 and 30 rapid amplification of cDNA ends (RACE) has been used with primers derived from regions of the EcR cDNA shared among all the isoforms. These extensions from a common region within the EcR sequence such as the DBD or LBD permit the rapid recovery of novel transcripts with isoform-specific coding regions (e.g., Minakuchi et al., 2002). The recovery of the D. melanogaster ultraspiracle (usp) gene resulted from three independent screens using vertebrate RXR as a probe (Oro et al., 1990),

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Figure 4 Amino acid alignment of representative insect EcR sequences (order Diptera: D. melanogaster, Lucilia cuprina, Aedes aegypti, C. tentans; order Lepidoptera: Heliothis virescens, Bombyx mori, Manduca sexta, Choristoneura fumiferana; order Orthoptera: Locusta migratoria; order Coleoptera: Tenebrio molitor). Also included: ixodid tick (Ambyomma americanum), and rat FXR. Amino acids designate number of first residue in alignment. References indicate original report. Bars designate each of the 12 helices, and shaded residues indicate those which have been mutated and reported (references given in the text). Binding sites of HvEcR with 20E indicated with circle (o).

Figure 5 Continued

252 The Ecdysteroid Receptor

employing an oligonucleotide derived from a highly conserved portion of the DBD to screen an embryonic cDNA library (Henrich et al., 1990), and by using a promoter element in the s15 chorion gene to screen an expression library (Shea et al., 1990). A genomic clone residing near the tip of the D. melanogaster X chromosome rescues larvae homozygous for the early larval lethal mutation, ultraspiracle (usp; Oro et al., 1990), so named because mutant larvae fail to shed their first instar cuticle and appear to develop an extra row of spiracles (Perrimon et al., 1985). The same combination of PCR-based and library screening approaches used for finding EcR have led to the recovery of complete USP sequences from other insects. The discovery that E75 was not only a target of ecdysteroid action, but a nuclear receptor itself, proved to be the first of several serendipitous discoveries of other nuclear receptors in the Drosophila genome that are associated directly with the salivary gland puffing response mediated by EcR and USP. Another ‘‘early late’’ puff gene product, E78, is a member of the nuclear receptor superfamily (Stone and Thummel, 1993) as is DHR3, which had been recovered in the same library screen that yielded EcR (Koelle et al., 1992); DHR3 is also a heterodimeric partner of EcR (White et al., 1997). Still another nuclear receptor, bFTZ-F1, a transcriptional regulator of the embryonic segmentation gene fushi tarazu (Lavorgna et al., 1991) is the product of the stage-specific 75CD chromosomal puff that occurs during the mid-prepupal ecdysteroid peak (Woodard et al., 1994). More systematic screens have yielded other Drosophila hormone receptors (DHRs), including DHR4 (Sullivan and Thummel, 2003), DHR39 (Horner et al., 1995), and DHR78 (Fisk and Thummel, 1998), which are orphans and ecdysteroid-regulated. Based on USP’s possible homology to RXR, a yeast two-hybrid assay employing USP as bait led to the isolation of an interacting orphan receptor, DHR38, that is orthologous to the vertebrate nerve growth factor 1B (NGF1B; Sutherland et al., 1995). The seven-up (svp) gene, originally defined by a mutation that disrupts photoreceptor function, encodes the Drosophila ortholog of another vertebrate orphan, the chicken ovalbumin upstream promoter-transcription factor (COUP-TF; Mlodzik et al.,

1990). COUP-TF heterodimerizes with RXR, and analogously, SVP forms a functional heterodimer with USP as shown by its ability to compete for dimerization with EcR in both D. melanogaster (Zelhof et al., 1995a) and A. aegypti (Zhu et al., 2003a). Therefore, the orphan nuclear receptors not only resemble EcR and USP structurally, it is increasingly apparent that they also play a role in the orchestration of ecdysteroid response, both as targets of EcR and USP-mediated transcriptional activity and as heterodimeric partners for the two functional components of the ecdysteroid receptor. Each of these orphans is sufficiently unique in sequence that a combination of library screening and PCR approaches have led to the successful recovery of orphan orthologs. A listing of the orphans implicated in some aspect of ecdysteroid regulation from other insect species is given in Table 1. Later in the chapter, the regulation and developmental roles of these orphans in D. melanogaster and other insects will be explored and compared further. The transcript levels of all 21 nuclear receptors found in the D. melanogaster genome have been analyzed preliminarily to reveal their potential connection with ecdysteroid-regulated developmental events. Some of these have not been functionally associated with ecdysteroid action and play a role in other developmental processes (Sullivan and Thummel, 2003, and references therein); these will not be considered further in this chapter. 3.5.2.3. Homology Models and Crystal Structure for Ecdysteroid Receptor Components

The need to understand the interaction of EcR with its natural ligand and other ecdysteroid agonists has prompted efforts to elucidate the crystal structure of the EcR LBD. Homology models based on a comparison of the EcR LBD with the known crystal structures of the vertebrate retinoic acid receptor (RAR) and vitamin D receptor (VDR) have been employed to formulate predictions about the threedimensional structure of the EcR LBD (Wurtz et al., 2000). The docking model resulting from this comparison with known crystal structures suggests that the ligand-binding pocket of EcR consists of a shallow tube that is just large enough to accommodate 20E and a bulky envelope. The shape and size of the

Figure 5 Amino acid alignment of representative insect USP sequences (order Diptera: D. melanogaster, L. cuprina, A. aegypti, C. tentans; order Lepidoptera: H. virescens, B. mori, M. sexta, C. fumiferana; order Orthoptera: L. migratoria; order Coleoptera: T. molitor ). Also included: ixodid tick (A. americanum) and human RXR-a. Amino acids designate number of first residue in alignment. References indicate original report. Bars designate each of the 12 helices, and shaded residues indicate those which have been mutated and reported (references given in the text).

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t0005

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Table 1 Nuclear receptors experimentally associated with ecdysteroid action in Drosophila melanogaster. Relevant references given in text Drosophila receptor

Noninsect ortholog

Insect orthologs

Mutational effect in Drosophila

EcR USP DHR4 DHR38 DHR39 E75

vert. FXR vert. RXR

see Figure 4 see Figure 5

vert. NGF1B

Manduca Aedes

Embryonic/larval lethal Larval lethal Not reported Disrupted cuticle development No discernible effect Larval/adult lethal

Metapanaeus (shrimp)

E78

Vert. PPARg Dirofilaria (filarial parasite)

DHR3

C. elegans CHR3, ROR

DHR78 FTZF1 SVP

vert. Steroidogenic factor-1 (SF1) vert. COUP-TF

Aedes, Bombyx, Choristoneura, Galleria, Manduca

Aedes, Choristoneura Galleria, Manduca, Bombyx, Tenebrio Bombyx, Tenebrio Aedes, Bombyx, Manduca, Tenebrio Aedes, Tenebrio

pocket predicted by this model also indicate that the more compact bisacylhydrazines that behave as nonsteroidal agonists of EcR fill up only a portion of the pocket. The crystal structure of HvEcR has shown that the EcR LBD is highly flexible, and that the shape of its ligand-binding pocket adapts to the ligand that occupies it, whether it be 20E, or a nonsteroidal agonist (Figure 6; Billas et al., 2003). Just as the crystal structure predicts, substitution of an amino acid that normally contacts 20E (A398P in HvEcR; see Figure 4 for its position in other EcR sequences) destroys HvEcR’s ability to mediate an inducible transcriptional response to 20E, but the same mutation does not affect the inducibility caused by a nonsteroidal agonist. The crystal structure further predicts that a valine residue residing in helix 5 of Lepidoptera EcR sequences (V384 in HvEcR) is responsible for the high affinity of a lepidopteran-specific agonist, BY106830. Most insect EcRs from other orders encode a methionine at this position in helix 5 (see Figure 4). The flexibility of the EcR LBD which allows it to bind structurally disparate ligands, presumably is stabilized by heterodimerization with USP. When USP is purified for crystal structure analysis, a phospholipid is copurified that is partially embedded in the relatively large ligand-binding pocket of USP. As a consequence, the USP ligandbinding domain is held in an antagonistic, apoconformation in which the carboxy-terminal region including the last alpha-helix (helix 12) is held out (Figure 7; Billas et al., 2001; Clayton et al., 2001). Another modeling study has shown that the USP LBD possesses the theoretical capability to reconfigure itself into an agonist position and interact with protein cofactors, although the residues involved in

Disrupted chromosome puffing Embryonic lethal Larval lethal Embryonic/larval lethal Disrupted eye development

these interactions would be different from those exposed on the RXR-a homolog when it assumes an agonist conformation (Sasorith et al., 2002). Modelling has also been employed to determine the capability for the three major juvenile hormones (JH1, JH2, and JH3) to bind to the ligand-binding pocket of USP. At least two ligand-binding configurations were identified, one associated with USP forms that carry an arginine residue at a critical position in helix 5 (see Figure 5). This residue prevails among the non-dipteran species for which the model predicted that JH acids could form a stable binding configuration. For the dipteran species, which typically carry a nonpolar amino acid at this position in helix 5, JH esters showed better affinity than JH acids, consistent with the results of biochemical binding studies performed on the Drosophila USP in which a purified USP fusion protein shows half-maximal binding in the range of 1–5 mM using a tryptophan fluorescence assay (Jones et al., 2001). While the modeling demonstrates the theoretical capability for various JH forms to bind to either lepidopteran or dipteran forms of USP, the occupancy rate of the ligand-binding pocket for them is relatively low for nuclear receptors. Nevertheless, other studies indicate that biological relevance does not always depend upon a high ligand binding affinity (Forman et al., 1995; Kitareewan et al., 1996; Staudinger et al., 2001). For instance, the affinity of the vertebrate peroxisome proliferator activating receptors (PPARs) involves ligand interactions of low specificity and affinity that approximates the levels predicted for USP and JH forms (Schmidt et al., 1992). Viewed from this perspective, the low affinity of a receptor for its activating ligand actually suggests a

Figure 6 Two ligand binding modes for HvEcR LBD as described in Billas et al. (2003). (a) Two weighted omit stereoview maps of the electron density for ponasterone A-bound HvEcR-LBD at 2.9 A˚ resolution. Ligand shown in blue and selected amino acid residues shown in magenta. Interactions between residues and ligand indicated as green dotted lines. (b) Schematic representation of the interactions between ligand-binding residues and ponasterone A. Arrows correspond to hydrogen bonds between ligand and amino acid residues. Residues in blue are common to both stereoview structures in (a), residues in white are those depicted only in the first stereoview map, and those in yellow are depicted only in the second (righthand) map. (c) two weighted stereoview maps, as above, with the nonsteroidal agonist, BYI06830. (d) schematic representation of the interactions between ligand-binding residues and BYI06830. (Adapted with permission from Billas, I.M., Iwema, T., Garnier, J-M., Mitschier, A., Rochel, N., et al., 2003. Structural adaptability in the ligand-binding pocket of the ecdysone hormone receptor. Nature 426, 91–96.)

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Figure 7 The dimerization structures of HvEcR LBD and HvUSP LBD. (a) EcR/USP LBD heterodimer with bound ponasterone A. (b) EcR/USP LBD heterodimer with nonsteroidal agonist, BYI06830. Helix 12 is shown in red, USP shown in blue. The missing loop that connects USP Helix 5 to the b-sheet is shown as a dashed blue line that projects closely to EcR helix 9–10. (Adapted with permission from Billas, I.M., Iwema, T., Garnier, J-M., Mitschier, A., Rochel, N., et al., 2003. Structural adaptability in the ligandbinding pocket of the ecdysone hormone receptor. Nature 426, 91–96.)

Figure 8 An idealized palindromic repeat element with a single nucleotide spacer used in the EcR-USP complex. Symbols as in Figure 3. Red indicates USP interaction sites, blue indicates EcR interaction sites. Solid arrows indicate a direct base interaction and hollow arrows indicate a water-mediated base interaction. Solid bars indicate a direct phosphate interaction, and hollow bars indicate a water-mediated phosphate interaction. Circles indicate van der Waals interaction. (Reproduced with permission from Devarakonda, S., Harp, J.M., Kim, Y., Ozyhar, A., Rastinejad, F., 2003. Structure of the heterodimeric ecdysone receptor DNA-binding complex. EMBO J. 22, 5827–5840.)

mechanism by which fairly tight regulation is achieved when ligand titers reach relatively high levels in the cell. A discussion of functional experiments concerning the effects of JH on the ecdysteroid receptor components, EcR and USP, is presented later. The crystal structure of the EcR and USP DBD at a canonical EcRE DNA-binding site reveals the nature and location of DNA-binding residues for the two DBD regions, as well as their points of proteinprotein interaction (Figure 8). While RXR forms a functional dimer with EcR (Christopherson et al., 1992; Yao et al., 1993), the physical interaction

between USP/RXR and EcR is different than it is for RXR with several of its natural partners (Figure 9; Devarakonda et al., 2003). The ability of the DHR38/USP heterodimer to induce transcription in response to ecdysteroids, even though the complex does not physically interact with its ligand, has prompted a crystal structure analysis of the DHR38 LBD. DHR38 lacks a true ligand-binding pocket – the space is filled with four phenylalanine side chains. The dimerization interface apparently lies along helix 10, but DHR38 lacks helix 12, and its AF2 function may reside in an unique sequence lying between helix 9 and 10 in an activated (agonistic) conformation (Baker et al., 2003). 3.5.2.4. Biochemical Analysis of EcR and USP

Numerous biochemical experiments have been performed to assess the properties associated with EcR and USP function in insects, including: (1) the affinity of EcR for ecdysteroids and nonsteroidal agonists, (2) the ability of EcR and USP to dimerize and interact with DNA target sequences, (3) the physical interaction of EcR and USP with other orphan receptors and proteins, and (4) the detection and demonstration of posttranslational and other modified forms of EcR and USP. The necessity for heterodimerization between EcR and USP to produce a functional ecdysteroid receptor was demonstrated by showing that the in vitro translated EcR and USP products bind to a radiolabelled hsp27 EcRE, but that neither product alone is capable of binding to the same element, based on the results of an electrophoretic mobility shift assay (EMSA). Moreover, this effect is enhanced by the simultaneous presence of muristerone A or

256 The Ecdysteroid Receptor

p9005

20E, indicating that the hormone stabilizes the heterodimer at an EcRE site. Further, the affinity of radiolabelled 125I-iodoponasterone, an ecdysteroid with high specific activity and affinity for EcR (Cherbas et al., 1988), is substantially higher for EcR/USP dimers than for EcR alone (Kd ¼ 1 nM; Yao et al., 1993). A similar level of ligand affinity is obtained with extracts taken from cultured Drosophila Kc and S2 cells, which contain endogenously expressed USP (Koelle et al., 1991). EcR also heterodimerizes with the vertebrate RXR and this heterologous dimer is responsive to muristerone A, but not 20E, indicating that USP plays some role in determining the ligand specificity of the complex (Christopherson et al., 1992; Yao et al., 1993). The ability of EcR and USP candidates to heterodimerize on an hsp27 EcRE has become a standard for verification (e.g., Swevers et al., 1996; Hannan and Hill, 2001; Durica et al., 2002; Minakuchi et al., 2002). The EcR/USP heterodimers from other insects show an affinity for 125I-iodoponasterone and [3H]ponasterone A that approximates the DmEcR/USP heterodimer (e.g., Kd ¼ 1.1 nM; Swevers et al., 1996). For AaEcR and AaUSP, the detection of EcR/USP heterodimers in a mammalian cell culture extract increases approximately 25-fold with the addition of 5 mM 20E. This accompanies a decrease in the quantity of AaUSP/AaSVP heterodimers, thus illustrating the competitive dynamics of these two protein–protein interactions involving USP (Zhu et al., 2003a). The ability of nonsteroidal agonists, such as the diacylhydrazines, RH5849 (1,2-dibenzoyl-1-tertbutylhydrazine) and RH5992 (tebufenozide), to displace radiolabelled ponasterone has been used to assess EcR specificity for these compounds. When extracts from a lepidopteran (Sf9) and dipteran insect cell line (Kc) are compared for the ability of the diacylhydrazines to displace [3H]-ponasterone A, the affinity of all RH compounds is considerably lower in the Kc cells (Nakagawa et al., 2002). Also, none of several RH compounds fail to displace [3H]-ponasterone A in extracts containing the migratory locust’s EcR and USP, indicating that the orthopteran EcR, like those of other ancient insect orders, possesses substantially different binding properties than those of the Lepidoptera (Hayward et al., 2003). Ligand binding requires the entire LBD, and deletion of a portion of helix 12 at the carboxy-terminal end of EcR is sufficient to eliminate ponasterone A binding (Perera et al., 1999b; Hu et al., 2003; Grebe et al., 2003). Fusion proteins expressing wild-type and mutated versions of DmEcR and DmUSP LBDs in yeast cells have also been subjected to tests of their affinity

for ponasterone A in cell extracts (Grebe et al., 2003). The association rate of a DmEcR LBD fusion protein is modestly elevated by the presence of USP, and the dissociation rate is reduced by about 20fold, so that the dissociation constant is reduced from about 40 nM in EcR alone to about 0.5 nM in the presence of USP. At least one mutation of the Drosophila EcR LBD (N626K) alters the rate of dissociation without affecting association rate, suggesting that an ecdysteroid enters and exits the EcR ligand-binding pocket by different routes, though this effect has not been tested in the intact EcR or in vivo. Purified DmUSP has the potential to form homodimers and other oligomers, even in the absence of a DNA-binding site, though this capability is reduced by removal of the A/B domain (Rymarczyk et al., 2003). Along with its ligand-binding properties, the ability to recognize a DNA element, usually the consensus hsp27 EcRE by an EcR/USP or EcR/RXR dimer, is a standard for identifying a functional ecdysteroid receptor. The hsp27 EcRE was originally localized by DNAse I footprinting and its ability to confer ecdysteroid-inducibility on an otherwise noninducible promoter in S1 Drosophila cultured cells. The sequence is 23 bp long and arranged in an inverted palindrome separated by a single nucleotide (Riddihough and Pelham, 1987). The EcR and USP zinc finger interactions with an idealized palindromic EcRE and with each other are described diagrammatically (Figure 3; Ozyhar and Pongs, 1993; Devarakonda et al., 2003) and shown graphically (Figure 8). A diagrammatic depiction of the nature of individual nucleotide interactions between the element and the zinc fingers is also shown in Figure 9 (Devarakonda et al., 2003). A closely related response element taken from an ecdysteroid-inducible gene in Kc cells, Eip28/29, has also been used experimentally (Cherbas et al., 1991). The EcR/USP heterodimer also recognizes half sites arranged into direct repeats (DR) separated by 0–5 nucleotide spacers (DR0–DR5) with the DR4 element showing the highest affinity. Various direct repeat elements were tested with the ecdysteroid-inducible and fat body-specific fbp1 promoter. When these direct repeat elements are connected to the minimal fbp1 promoter, they are unable to induce higher transcription in the presence of ecdysteroids. However, when the DR0 and DR3 elements are substituted for the natural fbp1 EcRE in its normal promoter context, both elements are ecdysteroid-inducible and fat body specific (Antoniewski et al., 1996). Context is also important for the intermolt, sgs4 gene promoter, which requires that an EcRE be surrounded by serum element binding protein

p9010

The Ecdysteroid Receptor

Figure 9 A superposition of the DmEcR/DmUSP/IR1 DNA complex and the DmEcR/HsRXR/IR1 DNA complex. USP is shown in red, RXR is shown in gold. EcR with USP is shown in purple, and EcR with RXR is shown in light blue. Zinc ions are shown in green and the single base pair spacer nucleotide is shown as magenta. (Reproduced with permission from Devarakonda, S., Harp, J.M., Kim, Y., Ozyhar, A., Rastinejad, F., 2003. Structure of the heterodimeric ecdysone receptor DNA-binding complex. EMBO J. 22, 5827–5840.)

(SEBP) binding sites in order to mediate a receptorinducible response (Lehman and Korge, 1995). A perfect palindrome carrying the half-site sequence, GAGGTCA separated by a single A/T nucleotide (PAL1) shows the highest affinity for the DmEcR/DmUSP complex in extracts taken from Drosophila S2 cells. By testing the ability of other elements to compete for this element, the order of affinity is been determined: PAL1 > DR4 > DR5 > PAL0 > DR2 > DR1 > hsp27,DR3 > DR0. In all cases, affinity is elevated by the presence of muristerone A. In cell culture experiments testing the inducibility of these elements when attached to a minimal promoter, inducibility correlates with affinity, except that the hsp27 EcRE is a stronger inducing element than DR1, DR2, or PAL0. When DR4 is placed in a reverse orientation relative to the promoter, it maintains its ability to mediate a transcriptional response (Vogtli et al., 1998). A similar correlation between DNA affinity and inducibility has been noted for the AaEcR/AaUSP complex and its response elements (Wang et al., 1998). A natural direct repeat EcRE has been identified in the ecdysteroid-regulated intermolt 3C puff of

257

D. melanogaster that is separated by 12 nucleotides (DR12) which is capable of conferring ecdysteroidresponsiveness to a heterologous promoter in cell cultures (D’Avino et al., 1995). The affinity of the EcR/USP complex for the DR12 element is offset by the presence of other orphan receptors (DHR38, DHR39, bFTZF1), and a weak inductive response to 20E is also offset by the presence of these orphans (Crispi et al., 1998). A similar competition scenario exists between DmEcR and DmSVP as heterodimeric partners with DmUSP. The SVP/USP dimer preferentially binds to a DR1 element, whereas the EcR/USP dimer displays a heightened affinity for an Eip28/29 EcRE (Zelhof et al., 1995a). The recognition of these canonical elements also depends upon specific features of the ecdysteroid receptor itself. In at least one instance, DNA recognition depends upon isoform-specific EcR and USP combinations. The EcR heterodimers formed with each of the two USP/RXR isoforms in the ixodid tick, A. americanum, display substantially different affinities for DNA response element sequences, with an RXR1/EcR dimer showing strong affinity for both palindromic and direct repeat sequences, whereas an RXR2/EcR dimer shows only weak affinity for a PAL1 element (Palmer et al., 2002). Ligand-binding specificity also plays a role in determining the affinity of EcR/USP for a response element. AaEcR/DmUSP recognition of an hsp27 EcRE on EMSAs is normally enhanced by the presence of ecdysone, but only 20E enhances affinity of the DmEcR in this assay. Domain swapping between the AaEcR and the DmEcR localizes a region within the LBD that is responsible for the differential effect of ecdysone on the two EcRs. Within the swapped region, a single amino acid conversion in the DmEcR (Y611F) to the corresponding AaEcR sequence results in a mutated receptor that shows the ability to dimerize in the presence of both 20E and ecdysone (Wang et al., 2000). Interestingly, all insect EcR sequences encode either a tyrosine or phenylalanine at this position in helix 10, which lies along a dimerization interface. The subtle yet significant difference in sequence, when considered in the framework of the numerous residue differences that exist among the insect EcR LBDs, illustrates the dimensions of potential functional diversity among them. Generally, the interpretation of EMSA results must be handled circumspectively because purified EcR/USP proteins do not always exhibit the same DNA binding properties as cell extracts which include the receptor components (e.g., Lan et al., 1999; Palmer et al., 2002). This is exemplified in the extreme by the observation that affinity column

258 The Ecdysteroid Receptor

purified DmEcR and DmUSP, when mixed together, fail to bind to the hsp27 EcRE unless other chaperone proteins are added. While no single chaperone is essential for DNA binding, the elimination of any one of them proportionally reduces EcRE recognition. The formation of this chaperone/receptor complex also requires ATP, although none of these chaperones is necessary for ligand binding. Only two of the six chaperones described, Hsp90 and Hsc70, are Drosophila proteins; the others used in the study were human chaperone proteins (Arbeitman and Hogness, 2000). By expressing GST fusion proteins in E. coli and using affinity chromatography to purify them, preparations of Chironomus EcR and USP have been recovered which retain essential DNA-binding and ligand-binding characteristics as long as detergents are absent from the purification process. Scatchard plots of bacterially expressed CtEcR reveal two high-affinity ecdysteroid binding sites, as do extracts from a Chironomus epithelial cell line (Grebe et al., 2000). Bacterial GST-fusion proteins expressing the Drosophila EcR and USP DNA-binding domains have also been isolated for the purpose of testing their affinity to DNA elements, although the GST motif alters the DNA-binding properties of the fusion protein (Niedziela-Majka et al., 1998; Grebe and Spindler-Barth, 2002). Similarly, the recovery of a fusion protein encoding the DmEcR LBD is enhanced by cleaving the C-terminal F-domain, which is considerably longer in D. melanogaster than other species (Halling et al., 1999); this cleavage exerts no discernible effect on transcriptional activity (Hu et al., 2003). Since EcR and USP function as part of a protein complex, orphan receptors and cofactors described throughout the chapter typically require the discovery and/or demonstration of a physical interaction. These demonstrations include the use of yeast two-hybrid assays (e.g., Sutherland et al., 1995; Beckstead et al., 2001), the interaction of in vitro translated EcR and USP with a protein or protein domain (e.g., Bai et al., 2000), and the identification of interacting proteins that coprecipitate with EcR and/or USP (e.g., Sedkov et al., 2003). A member of another class of proteins, the immunophilins, which are known to interact with vertebrate steroid receptor complexes, coprecipitates as part of an EcR/USP complex taken from M. sexta prothoracic (ecdysteroidogenic) glands. Ligand affinity of the complex falls within an expected range, though it is unknown whether the interaction between the immunophilin, FKBP46, and EcR/USP is direct and also unknown whether this interaction affects transcriptional activity (Song et al., 1997).

The colocalization of proteins such as RNA polymerase II and EcR on Chironomus polytene chromosomes by immunostaining illustrates a possible relationship between the complex and the cell’s transcriptional machinery (see Figure 1; Yao et al., 1993; Wegmann et al., 1995). A zinc finger protein that preferentially localizes at active puffs along Drosophila polytene chromosomes, known as peptide on ecdysone puffs (PEP), has been characterized by anti-PEP antibody decoration of polytene chromosomes, but has not yet been functionally associated with ecdysteroid action (Amero et al., 1991). The effect of posttranslational modifications upon ecdysteroid activity has also not been explored extensively, though a phosphorylated form of DmUSP in fly larvae has been identified by showing that phosphatase activity eliminates a high molecular weight band in larval extracts. The phosphorylated form of USP in larval salivary glands is elevated by incubation with 20E (Song et al., 2003). Similarly, the presence of a phosphorylated form of both CtEcR and CtUSP extracted from C. tentans larvae has been demonstrated on Western immunoblots (Rauch et al., 1998). The relative abundance of phosphorylated forms of the two USP isoforms in the M. sexta prothoracic gland is altered by the presence of 20E and has been associated with changes in ecdysteroidogenic activity, suggesting a feedback mechanism by which ecdysteroid synthesis is downregulated in prothoracic glands (Song and Gilbert, 1998). While several groups have employed DNA cloning methods to recover receptor genes from which protein sequences are deduced, other efforts have demonstrated receptor purification through measurements of their activity. Partial purification of an ecdysteroid-binding protein from Drosophila embryos produces two peptides that are radiolabelled by a 20E derivative, one of 150 kDa and another of 90 kDa (Strangmann-Diekmann et al., 1990), the latter being the molecular weight predicted from the Drosophila EcR cDNA sequence (Koelle et al., 1991). Purification of the protein through its affinity to a magnetic hsp27 EcRE failed to resolve the disparity between the measured size of the purified product and the predicted size of the protein (Ozyhar et al., 1992). Purification of the 20E receptor yielded a protein that binds to the appropriate response element sequence of the hsp27 promoter, as well as an hsp23 promoter element, and elevates transcription rates by 100-fold (Luo et al., 1991). The presence of ecdysteroids increases the affinity of the receptor protein as measured by EMSA.

p0230

The Ecdysteroid Receptor

3.5.3. Functional Characterization of the Ecdysteroid Receptor 3.5.3.1. Cell Culture Studies: Rationale

The study of EcR and USP in vivo is complicated because ecdysteroid responses vary both spatially and temporally. In fact, the effects of ecdysteroids can be obscured by the heterogeneity of response at the promoter of a single gene (Andres and Cherbas, 1992; Andres and Cherbas, 1994). Therefore, it is often difficult to unravel in vivo transcriptional and cellular responses mechanistically, even in a single cell type, since the response to ecdysteroids typically triggers ongoing changes in the target cell, that in turn, affect later ecdysteroid responses. Cell cultures provide the benefit of working with a stable, relatively homogeneous cell type that does not require the rigorous staging and rearing conditions necessary for meaningful studies in vivo. Thus, cultured cells can provide a variety of important and useful insights about EcR and USP by establishing a foundation for subsequent in vivo hypothesis testing. Because they are easy to grow, it is also relatively easy to recover cellular extracts for biochemical testing. Nevertheless, stably cultured cells probably do not duplicate any actual cell exactly, so that observations reflect what a cell can do, but not necessarily what a cell does. Therefore, subsequent in vivo verification is essential for insights garnered through cell culture experiments. Numerous experiments involving cultured cell lines have been conducted over the years to test the ability of the ecdysteroid receptor to regulate transcriptional activity induced by ecdysteroid treatment. Early experiments focused on endogenous changes in cell morphology and gene expression in insect cells challenged with ecdysteroids such as 20E, which express EcR and USP endogenously. As transfection technology has developed, increasingly sophisticated cell systems have been developed to analyze and dissect receptor functions. In recent years these systems have been developed to screen for novel agonists and devise ‘‘new and improved’’ ecdysteroid induction systems for agricultural, biomedical, and commercial application (see Chapter 3.4). 3.5.3.2. Cell Cultures: Characterization

The most prominent cell line for early studies of ecdysteroid action was the D. melanogaster Kc cell line, and derivatives of it are still employed today (Hu et al., 2003). In response to 20E at 107 to 108 M, Kc cells start to develop extensions within hours, produce acetylcholinesterase, and undergo a

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proliferative arrest a few days later. These cells are even more responsive to the phytoecdysteroids, ponasterone A and muristerone A, than to 20E (Cherbas et al., 1980 and references therein). With prolonged exposure to ecdysteroids, Kc cells become insensitive to 20E and EcR levels become depressed (Koelle et al., 1991). The nonsteroidal agonist RH5849 not only mimics the effects of 20E when tested on sensitive Kc cells, it also fails to evoke a response from insensitive Kc cells, thus arguing that the RH5849 acts via a common mechanism with ecdysteroids (Wing, 1988). Other cell lines, Schneider 2 and 3 (S2 and S3), also become insensitive to ecdysteroid action by reducing their titer of EcR, and have been used because they are relatively easy to transfect transiently with fusion and reporter genes. As noted earlier, the identity of the DmEcR gene was partly demonstrated by introducing the DmEcR cDNA into ecdysteroid-insensitive S2 cells and thus restoring their responsiveness to 20E (Koelle et al., 1991). More recently, Kc cell lines have been improved by developing protocols for stable integration of transgenes by P-element transposition into the genome (Segal et al., 1996) and by using parahomologous gene targeting to ‘‘knock out’’ an endogenous gene target (Cherbas and Cherbas, 1997). In this way, a cell line, L57-3-11 containing no endogenous EcR has been produced which allows the introduction of modified versions of EcR for subsequent experimentation, without the complications normally posed by the presence of endogenous EcR (Hu et al., 2003). The Kc cell line has been used to identify ecdysteroid-regulated transcriptional targets. By comparing hormone-treated and control Kc cells, Savakis et al. (1980) isolated and identified three ecdysteroid-inducible peptides (EIPs) named by their molecular mass in kiloDaltons, EIP28, EIP29, and EIP40, whose synthesis is elevated 10-fold at 108 M 20E. JH supplementation of the cell culture medium inhibits acetylcholinesterase induction but not EIP induction (Cherbas et al., 1989). The use of ecdysteroid-responsive lines for other insects is expanding continuously, largely in order to understand the causes of insecticidal resistance to ecdysteroid agonists and to identify novel insecticidal candidates. A C. tentans epithelial cell line was used to isolate several stable and ecdysteroidresistant clones that metabolizes 20E relatively quickly (Spindler-Barth and Spindler, 1998). Clones from the same line were later selected for their resistance to the inductive effects of RH5992 and differential metabolism of RH5992 was ruled out as a cause for the resistance. While the profile of EcR in these cells is approximately normal, some

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lines shows a modest decline in the abundance of phosphorylated USP forms that is not reversible by addition of 20E. A variety of differences in EcR ligand-binding characteristics were noted among the cell lines tested that presumably reflect the role of unknown factors underlying the resistance (Grebe et al., 2000). Another widely used lepidopteran cell line is Sf9, derived from ovarian cells of the fall armyworm, Spodoptera frugiperda. The affinity of ponasterone A for Sf9 and Kc cells is very similar, but the nonsteroidal agonist ANS-118 (chromafenozide) displays a much higher affinity for extracts derived from Sf9 cells than from Kc cells (Toya et al., 2002). Similarly, cell lines derived from C. fumeriferana show a much greater responsiveness to RH5992 than Drosophila cultured cells, which do not respond to the compound and excrete it an elevated rate via an ABC transporter system (Retnakaran et al., 2001). Choristoneura cell lines, with prolonged exposure to RH5992 become irreversibly insensitive to ecdysteroids and to the agonist, though the mechanism is unknown (Hu et al., 2004). A more comprehensive discussion of ecdysteroid agonist properties is provided elsewhere in this series (see Chapter 3.4), though it is clear that the differences among species are derived primarily from structural diversifications in EcR and possibly, USP (see Chapter 3.6). Other lines are particularly useful for studying ecdysteroid action in economically important insect species even when a complete EcR and USP have not been yet been isolated. These lines include the European corn borer (Ostrinia nubilalis; Trisyono et al., 2000), the tent caterpillar (Malacosoma disstria; Palli et al., 1995), the fall armyworm (Spodoptera frugiperda; Chen et al., 2002a), and the cotton boll weevil (Anthonomus grandis; Dhadialla and Tzertzinis, 1997). The ecdysteroid responsiveness has been established for at least some of these lines by showing that orthologs for ecdysteroid responsive D. melanogaster genes, notably the genes encoding E75 (early puff) and DHR3 (early-late puff), are activated transcriptionally when the cells are challenged with 20E or an agonist. A major benefit of using insect cell lines is that they provide the cofactors and machinery that are necessary for transcription to occur. On the other hand, other factors may modify or obscure an aspect of ecdysteroid response in a given experimental regime as silent and unidentified participants. As is evident from in vivo studies, a given experiment will generate substantially different cellular responses depending upon the milieu of proteins that contribute to ecdysteroid responsiveness in a given cell type. For this reason, mammalian cell lines containing no

endogenous ecdysteroid responsiveness have been used to reconstruct an ecdysteroid responsive transcriptional system by introducing the genes encoding EcR, USP, cofactors, and reporter genes in order to analyze their individual effects on ecdysteroid action (e.g., Christopherson et al., 1992; Yao et al., 1993; Palli et al., 2003; Henrich et al., 2003). Heterologous cells and cell lines also pose special difficulties, since it is conceivable that endogenous proteins provide novel functions or act as surrogates for similar insect proteins. Conversely, these cells might render false negative results in some experiments because they are missing one or more crucial cofactors. 3.5.3.3. Cell Cultures: Functional Receptor Studies

A variety of studies have examined the capability of EcR and USP to function in insect and other cell lines. Using the same approach, several orphan receptors from D. melanogaster have also been tested for their ability to respond to candidate ligands. These studies have generally focused on: (1) the responsiveness and activity of target DNA sequences in promoters, (2) the effect of various ecdysteroids, agonists, and antagonists upon receptor activity, (3) the properties of specific domains on receptor function and the effect of structural modifications upon transcriptional activity, and (4) the effect of heterodimerization and other cofactors, including orphan receptors, upon transcriptional activity. 3.5.3.3.1. Promoter studies The hsp27 EcRE connected to a weak constitutive promoter is capable of inducing the transcription of a reporter gene in a Drosophila cell line by over 100-fold when challenged with 20E (Riddihough and Pelham, 1987). When hsp27 EcREs are organized in a tandem repeat and attached to a weak promoter, the gene is not only responsive to 20E, but also is repressed in the absence of hormone, suggesting a repressive role for the ecdysteroid receptor via this element (Dobens et al., 1991). A plethora of cell culture transcriptional tests and accompanying EMSAs have been reported which followed from this early work, and this is an important criterion for determining the functionality of an EcR/USP heterodimer, as it was in the original report (Koelle et al., 1991). The most fundamental task concerning promoter activity is to define the functional promoter elements to which the ecdysteroid receptor complex binds and exerts its effects on transcription. This was accomplished for the genes encoding the aforementioned EIP28 and 29, which are translated from

The Ecdysteroid Receptor

alternatively spliced mRNAs of the same gene (Schulz et al., 1986). Three specific ecdysone response elements are ecdysteroid-responsive in vivo (Andres et al., 1992; Andres and Cherbas, 1994). One of these Eip28/29 EcREs lies in the promoter region though it is not required for ecdysteroidinducibility in Kc cells, possibly because this element is not used in the embryonic hemocytes from which the cells are derived. The other two elements lie downstream from the polyadenylation site, that is, on the 30 side of the gene (Cherbas et al., 1991). The EIP40 gene is also ecdysteroid-inducible in vivo (Andres and Cherbas, 1992) and the genomic region that includes the EIP40 gene has been analyzed (Rebers, 1999). Just as cell culture experiments have been used to identify EcREs in vitro for subsequent in vivo analysis, the reverse relationship has also been undertaken successfully – the ecdysteroid-responsive promoter elements in the b3 tubulin gene, which is ecdysteroid-responsive in vivo (Sobrier et al., 1989), were identified by testing their inducibility by 20E in Kc cells (Bruhat et al., 1993). Cell culture experiments have been used not only to dissect the functional ecdysteroid response elements within a promoter region, but also to examine the possibility that different factors compete for these sites to modulate transcriptional activity. The relationship of M. sexta EcRB1 and two MsUSP isoforms has been explored in M. sexta GV1 cell cultures by observing the ecdysteroid-inducible MHR3 promoter (Lan et al., 1997). Both EcR/USP complexes display about the same level of ligand affinity, but the EcRB1/USP1 complex induces much higher levels of expression in response to 20E from an intact promoter and also represses basal expression in the absence of 20E. Further, the EcRB1/USP1 complex binds to a canonical EcRE in the promoter and activates transcription, but the addition of USP2 prevents transcription and blocks binding to the EcRE. In vitro translated EcRB1/ USP2 can bind to this same EcRE, indicating that other cellular cofactors are responsible for the differential action of the two MsUSP isoforms (Lan et al., 1999). The two isoforms of CtUSP show somewhat different ligand-binding capabilities, and one of the forms, when purified, recognizes a DR1 element that is not recognized by the other. However, both show about the same ability to induce transcription with DmEcR in human HeLa cells. The CtEcR does not possess the ability to induce transcription in cell cultures (Vogtli et al., 1999). 3.5.3.3.2. Transactivation studies The appearance of multiple EcR isoforms in D. melanogaster has

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prompted comparisons of their transcriptional responsiveness in cell cultures and in yeast. The B1 and B2 N-terminal domains from DmEcR, when combined with a GAL4 DBD and and transfected into yeast, are capable of mediating transcription via a GAL4-responsive universal activation site (UAS) in the promoter. This AF1 (ligand-independent) transcriptional function is further elevated by removing an inhibitory region within the B1 domain (Mouillet et al., 2001). In human HeLa cells, DmEcRB1 and DmEcRB2 showed about the same level of AF1 function and both also possessed AF2 (ligand-dependent) transcriptional activity. By contrast, EcRA reduced basal transcriptional activity below the level obtained from an EcR with no A/B domain at all. Similar results have been obtained in a mammalian Chinese hamster ovary (CHO) line, except that the B1 isoform displays a highly elevated AF1 function compared to the A and B2 isoforms (Henrich et al., 2003). This activity difference measured for B1 in the two cell lines suggests that the HeLa cell line carries a repressor that interacts with the B1 domain which does not exist in CHO cells. It also illustrates the diversity of function that is possible among different cell types owing to differences in the cellular milieu. In the EcR-deficient L57-3-11 line, all three EcR isoforms are responsive to ecdysteroids, though the A domain contributes no AF1 function, nor does the DmUSP A/B domain. The B1 domain contains several regions that when deleted impose a modest impact on AF1 functions. By contrast, specific point mutations in the B2 domain dramatically reduce AF1 functions (Hu et al., 2003), suggesting that a localized interaction with other transcriptional factors is essential for activity. Lezzi et al. (2002) devised a yeast two-hybrid assay which fused the EcR and USP LBD to the yeast GAL4 activation domain and DNA-binding domain, respectively. A variety of site-directed mutations of the LBDs were used to test their effects on functional dimer formation and both AF1 and AF2 (ligand-dependent) transcriptional activity as measured by UAS-mediated lacZ activity. The wild-type LBDs dimerize and induce activity at a low rate even in the absence of muristerone A, and this rate increases more than tenfold in the presence of muristerone A (but not other ecdysteroids such as 20E). Mutations of critical residues in EcR’s helix 10 all but eliminate both dimerization and transcriptional activity, as expected. Deletion or mutation of helix 12, which normally folds over the ligand-filled pocket, eliminates EcR’s AF2 function, as do mutations that affect ligand binding, but AF1 function is still detected in these mutant forms of

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EcR. Substitution at a consensus cofactor-interacting residue, K497E, results in a strong elevation of basal transcriptional activity, though this mutant EcR has a reduced capacity for ligand-dependent induction and low ligand affinity (Bergman et al., 2004; Grebe et al., 2003). Virtually all point mutations in the DmUSP LBD eliminate transcriptional activity, though many retain the capability to dimerize with EcR, suggesting that USP is required for normal AF1 activity. Deletion of the carboxy-terminal region of DmUSP, including helix 12, does not disrupt homodimerization detected by gel filtration, but eliminates ligand-binding. Substitutions of specific residues in helix 12 modestly reduce basal transcriptional activity in yeast. However, while ligandbinding for the mutant USP is only slightly reduced, the ligand-dependent transcription of the EcR/USP complex is virtually eliminated (Przibilla et al., 2004). This result is counterintuitive since the crystallized DmUSP is locked into an antagonistic conformation which implies that AF2 activity is impossible. It also contrasts with the normal activity seen in several helix 12 mutations of USP that were tested in the Drosophila L57-3-11 cell line, which may express enough DmUSP to mask the mutational effects (Hu et al., 2003).

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3.5.3.3.3. Ligand effects At least three issues pertaining to ligand responsiveness of the ecdysteroid receptor have been addressed through the use of cell cultures. First, one has focused on the responsiveness of a given receptor to ecdysteroids and agonists, particularly novel compounds. Second, modified receptors have been tested for their capacity to respond to a given ligand or to assess the structural basis of ligand specificity. Finally, recent studies have focused on the possibility that other orphan receptors are responsive to ecdysteroids and/or other ligands. By testing ecdysteroid-induced transcriptional activity in cell culture, new nonsteroidal agonists, including both artificial compounds (Mikitani, 1996) and a variety of phytochemicals which act as agonists, including 8-O-acetylharpagide (Elbrecht et al., 1996) or antagonists, including several flavones (Oberdorster et al., 2001). Environmental chemicals including polyaromatic hydrocarbons (PAHs) and polychorinated biphenyls (PCBs) induce ecdysteroidresponsive genes, suggesting an EcR-mediated basis for molting defects commonly found among aquatic species residing in polluted water (Oberdorster et al., 1999). The use of cell culture assays has also been utilized to analyze the effects of juvenile hormone and its analogs upon transcriptional activity. The topic of juvenile hormone and its effects upon EcR and USP will be addressed later in the chapter.

Based on lines of evidence gleaned from experimental studies over the years, the possibility that ecdysteroids other than 20E display biological activity has been speculated about and even suspected for years (Somme-Martin et al., 1990; Grau and Lafont, 1994; Henrich and Brown, 1995). In S2 cells, the 20-hydroxylated derivates of the major and natural ecdysteroid products of the D. melanogaster larval ring gland, 20E and makisterone A are the most efficacious in activating a reporter gene whose promoter carried three tandem copies of the hsp27 EcRE. By contrast, precursor and metabolite ecdysteroids, as well as nonsteroidal agonists, evoke considerably less activity in the cell culture assay. As noted in other studies, ponasterone A displays the highest potency, and two other phytoecdysteroids, muristerone A and cyasterone, also are more inducible than 20E in these cells (Baker et al., 2000). Receptor modifications have also been utilized to delineate the basis for differences in ligand specificity, as noted earlier for the effects of the A398P mutation to test predictions based on the HvEcR crystal structure (Billas et al., 2003). Similarly, a substitution in the CfEcR (A393P) eliminates both its ligand-binding affinity for ponasterone A and its transcriptional inducibility in cell cultures, possibly by impairing the receptor’s interaction with a coactivator (GRIP1), but the mutation exerts no debilitative effect on responsiveness to a nonsteroidal agonist (Kumar et al., 2002). A chimeric Drosophila/Bombyx EcR carrying only the Bombyx D-domain is responsive to RH5992 in mammalian CV-1 cells. The effect is largely attributable to the relatively high level of RH5992-dependent dimerization between the D-domain of the chimera and endogenous RXR or cotransfected USP (Suhr et al., 1998). Based on a deletion analysis of EcR, both its D and E domain are required for ligand-dependent dimerization with USP (Perera et al., 1999b). As already noted, insect cells may provide endogenous components that influence ecdysteroid activity. EcR itself masks a second ecdysteroid responsive activity in Drosophila S2 cells. A fusion protein carrying the DHR38 LBD was tested for its ability to dimerize with DmUSP in response to several ecdysteroids using S2 cells from which endogenous EcR activity was eliminated by RNA interference. A response to 20E occurs that is further induced by adding an RXR activator, indicating that the DHR38/RXR heterodimer is responsible for the effect. Similarly, cotransfection with a chimeric and constitutively active form of USP also confers the ability of DHR38 to respond to 20E (Baker et al., 2003). The DHR38/USP dimer also responds to at least six other ecdysteroids, including some, such as

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3-dehydro-20E and 3-dehydromakisterone A, which are abundant during late larval development in Drosophila and display unusually high inducibility in fat body (Somme-Martin et al., 1990). The relevance of this pathway for ecdysteroid-driven events represents an important avenue for investigation in Drosophila and other insects. Cell culture experiments have also been used to demonstrate the effect of orphan receptors and receptor cofactors upon ecdysteroid response, including AHR38 (Zhu et al., 2000), AaSVP (Zhu et al., 2003a), DmSVP (Zelhof et al., 1995a), and DHR78 (Zelhof et al., 1995b). These will be discussed in connection with their biological relevance for modulating ecdysteroid receptor-mediated activities. 3.5.3.4. Ecdysteroid-Inducible Cell Systems

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The use of ecdysteroid-inducible cellular systems has emerged rapidly in recent years since ecdysteroids and nonsteroidal agonists are inexpensive and harmless to humans. Yeast assays have been developed to screen for ecdysteroid agonists of potential commercial value. When EcR by itself is transformed into yeast cells, it is capable of inducing a high level of transcription that is ligand-independent and which is not appreciably increased by the presence of USP or RXR (De la Cruz and Mak, 1997). Through modifications of the A/B domain, an inducible system was successfully produced (De la Cruz et al., 2000). A second ecdysteroid-inducible yeast system was produced by making at least two modifications. First, by removing the AaEcR A/B domain, the receptor’s AF1 function is reduced when transformed into yeast. Second, the addition of the mammalian receptor coactivator GRIP1 confers yeast cells with the ability to be induced more strongly by a range of ecdysteroid agonists. Conversely, the addition of the receptor corepressor SMRT represses transcription (Tran et al., 2001a). A yeast inducible assay utilizing the Cf EcR and Cf USP also requires the removal of the A/B domain from both EcR and USP, along with the addition of the GRIP1 coactivator (Tran et al., 2001b). In fact, the USP A/B domain often displays repressive properties or is inactive in heterologous cells, thus necessitating its deletion and/or replacement with a constitutively active N-terminal domain, such as VP16 (Tran et al., 2001b; Hu et al., 2003; Henrich et al., 2003). The field use of the bisacylhydrazines as insecticides has also inspired the introduction of ecdysteroid-inducible systems into plants which could be used to promote the expression of beneficial genes. Nonsteroidal agonist-inducible expression has been obtained from Zea mays (corn) protoplasts

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(Martinez et al., 1999b), tobacco, and Arabidopsis (Padidam et al., 2003) via EcR fusion proteins. This induction in Arabidopsis has been successfully linked to the expression of a coat protein gene from the tobacco mosaic virus (TMV), which confers resistance to TMV infection (Koo et al., 2004) and also induces the expression of a factor in maize that restores fertility to a male-sterile strain (Unger et al., 2002). As noted, ecdysteroids evoke no discernible responses upon mammalian cells, and therefore, attempts to implant an ecdysteroid-inducible gene expression system into cells for therapeutical purposes has been undertaken. EcR dimerizes with endogenously expressed RXR to form a functional heterodimer that responds to biologically inert ecdysteroids or nonsteroidal agonists (Christopherson et al., 1992; Palli et al., 2003). By producing and testing a variety of chimeras, a heterologous receptor system has been developed in mammalian cells that evokes induction rates of almost 9000-fold, with very rapid reduction when the ecdysteroid agonist is removed (Palli et al., 2003). A mouse strain carrying a transgenically introduced Drosophila EcR gene has been produced in which tissues are capable of responding to ecdysteroids without any other discernible phenotypes other than the expression of an hsp27 EcRE regulated reporter gene (No et al., 1996). The ecdysone switch has been shown to be responsive to several ecdysteroids and ecdysteroid agonists, and the maximal response can be further elevated by the addition of RXR activators. RXR superinduction requires that EcR be previously bound by its cognate ligand since RXR activators by themselves exert no effect upon EcR/RXR activity (Saez et al., 2000). Ecdysteroid-inducible expression of medically important genes has been accomplished in human cell lines (Choi et al., 2000), as well as the introduction of an ecdysteroid-inducible gene into rats via an adenovirus vector used for somatic gene therapy (Hoppe et al., 2000).

3.5.4. Cellular, Developmental, and Genetic Analysis 3.5.4.1. The Salivary Gland Hierarchy: A Model for Steroid Hormone Action

Many of the insights concerning steroid hormone regulation of transcriptional activity were first recognized in the larval salivary gland of several insect species. By observing the location, timing, and size of these chromosomal puffs, the activity of an ecdysteroid-inducible gene can be inferred. The progression of events, and their relation to

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developmental changes in the salivary gland and the whole animal are described here for D. melanogaster (see Thummel, 2002 and references therein). During the period of the third instar that precedes the wandering stage, ‘‘intermolt puffs’’ appear at several specific chromosomal sites. In response to the late larval ecdysteroid peak, these puffs regress and another set of ‘‘early’’ ecdysteroid-responsive puffs appear. The timing and duration of these ‘‘early’’ puffs varies (some are referred to as ‘‘earlylate’’ puffs) and these eventually regress as another set of ‘‘late puffs’’ appear at other chromosomal sites. The puffing pattern accompanies the production and secretion of glue protein from the salivary gland, which is extruded as the larva becomes immobile and tanning of the larval cuticle begins (puparium formation). Several hours later, a second smaller pulse of ecdysteroid induces a second wave of ‘‘early’’ puff activity, which is followed by a second wave of ‘‘late’’ puff activity. The two peaks evoke similar, but not identical, transcriptional changes, and these subtle differences underlie important developmental consequences. The second wave culminates in the expression of bFTZ-F1, a specific isoform of an orphan receptor encoded by the stage-specific (that is, the prepupal specific) gene puff located at the chromosomal interval, 75C. bFTZ-F1, in turn, sets off a cascade that ultimately includes the expression of another stage-specific puff, E93, involved in the histolysis of the gland (Broadus et al., 1999; Lee et al., 2002). As expected, 20E shows the greatest potency in terms of puff induction, followed by 3-dehydro20E, which is an endogenously produced ecdysteroid in the Drosophila third instar (Somme-Martin et al., 1988). Significantly, however, while the potency of individual ecdysteroids varies, no individual ecdysteroid evokes an unique puffing site, or alternatively, fails to evoke puffing at a site induced by other ecdysteroids. In other words, there is no evidence for multiple ecdysteroid signaling pathways in the Drosophila salivary gland (Richards, 1976). 3.5.4.2. Molecular and Genetic Characterization of the Puff Hierarchy

Numerous genetic and molecular studies have verified the basic tenets of the Ashburner model. Ashburner proposed that the ecdysteroid receptor directly induces the early puffs while simultaneously repressing late puff genes, and that the early puff gene products not only increase late puff transcription, but also feed back to downregulate their own expression. The use of chromosomal deletions and duplications of the region that contains E74 and

E75 provides a way to alter the intracellular dosage of early puff gene products and thereby test the model. Late puffs appear earlier and are larger when the early puff genes are duplicated, consistent with the accumulation of an activating factor. On the other hand, late puff appearance is delayed in chromosomes with early puff deletions (Walker and Ashburner, 1981). Further evidence that late puffing events are dependent upon early puffing events was shown ingeniously by testing permeabilized salivary glands 20E in combination with cellular extracts (Myohara and Okada, 1987). When ecdysteroid and a cellular extract from unstimulated salivary glands are mixed together, no response is registered from the late puff at 78C, but this puff appears when 20E, along with a cellular extract from stimulated salivary glands, is tested on permeabilized salivary glands. Another line of inquiry has focused on demonstrating that EcR and USP interact directly with puff site regions. Immunostaining has shown specific instances in which EcR and USP colocalize to early puff sites (Yao et al., 1993), and that EcR and RNA polymerase II antibodies colocalize on C. tentans polytene chromosomes (see Figure 1). EcR has also been associated with late puff sites (Talbot et al., 1993), inferring that it directly influences the regulation of late puff genes. Further, the products of the early puff gene, E75, have been associated by immunostaining with early and late puff sites, as predicted by the Ashburner model (Hill et al., 1993). The puff hierarchy actually begins before the onset of pupariation in Drosophila with the appearance of several intermolt puffs (see Lehman, 1995 and references therein). The sgs4 gene encodes a salivary glue protein and is regulated by the synergistic interplay of an ecdysone receptor element, and a secretion enhancer binding protein (SEBP3) site. A second SEBP binding site has been associated with products of the early puff product, Broad, indicating that the proper regulatory interplay between the ecdysone receptor and other transcriptional factors depends upon the cellular milieu (von Kalm et al., 1994; Lehman and Korge, 1995). As ecdysteroid titers increase, the intermolt puffs regress. Three early puff genes have been the subject of intensive ongoing study in D. melanogaster and other insects. All three genes encode transcription factors, consistent with a role in regulating late puff expression: the Broad-Complex (Br-C) encodes a zinc finger protein (DiBello et al., 1991), E74 is an ets protooncogene (Burtis et al., 1990), and E75 encodes an orphan receptor (Segraves and Hogness, 1990). All three possess a level of organizational

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complexity that is not evident from the cytological studies alone. Br-C encodes several different isoforms derived from alternative splicing (DiBello et al., 1991). Broad proteins carry a common core domain and a subset of several different variable domains (see Bayer et al., 1997), and the expression of each isoform varies both spatially and temporally (e.g., Huet et al., 1993; Mugat et al., 2000; Brennan et al., 2001; Riddiford et al., 2003). The switchover of expression among combinations of Br-C isoforms has been associated with the modulatory regulation of many ecdysteroid-inducible genes including the transcription factor gene hedgehog (hh; Brennan et al., 1998), micro-RNA (Sempere et al., 2003), and fbp1 (Mugat et al., 2000). The E74 puff encodes two isoforms, one of which is transcribed at low ecdysteroid levels (E74B; Karim and Thummel, 1991) and disappears as titers become elevated while the transcript which specifies the E74A isoform becomes more abundant. Finally, three isoforms exist for the nuclear receptor at the E75 puff (A, B, and C; Segraves and Hogness, 1990) resulting from alternative splicing. One of these, E75B, carries only the second zinc finger within its DBD. Several other ecdysteroid responsive puff genes have also been identified which play cellular roles (see Thummel 2002, and references therein). 3.5.4.3. Developmental Regulation of Ecdysteroid Response in D. melanogaster

Transcript levels of EcR, the intermolt genes, and the individual isoforms of the early puff genes (BR-C, E74, E75) vary temporally among individual tissues during the late larval/prepupal period in D. melanogaster (Huet et al., 1993). It is apparent that the relative abundance of the early puff isoforms changes continuously as ecdysteroid titers surge and decline, that the profile of ‘‘early puff’’ expression varies among tissues at any given time, and that the pattern changes are specific for a given tissue. For instance, EcR levels are very low throughout the larval/prepupal period in imaginal discs, whereas they remain elevated in the gut throughout this period. The relative abundance of EcR and its early puff products plays some role in the timing of subsequent gene expression, as exemplified by the Ddc (dopa decarboxylase) gene. It behaves as an ‘‘early late’’ gene and is induced by the combined action of EcR and Br-C, which is offset, in turn, by a repressive promoter element in epidermis and imaginal discs (Chen et al., 2002b). A similar mechanism has been described for the suppression of Ddc in M. sexta (Hiruma et al., 1995). The complexity of in vivo response to ecdysteroids is also revealed by comparing the puffing

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response seen along the polytene chromosomes of the larval fat body during the same time that the salivary gland puff response occurs (Richards, 1982). While many aspects of the fat body response resemble those seen in the salivary gland, individual fat body puff sites are responsive to both ecdysone and 20E in ranges below 106 M, whereas salivary gland puffing is much more responsive to 20E. While one obvious explanation lies with differences in metabolism (see Chapter 3.3), the fat body response to ecdysone does not lag the response to 20E in the fat body, as would be expected if conversion to 20E preceded receptor-mediated response. Fat body chromosomes also evoke different puffing patterns than salivary gland chromosomes, and these presumably reflect functional differences between the two compared cell types. Diversity of response has been seen for other ecdysteroid-inducible genes. For instance, the fat body protein-1 (fbp1) gene is ecdysteroid-inducible and expressed only in the fat body during the onset of metamorphosis (Maschat et al., 1991). The aforementioned Eip28/29 gene is ecdysteroid-responsive but displays several different patterns of expression in individual tissues at the onset of metamorphosis (Andres and Cherbas, 1992, 1994). The complexity of in vivo regulation is also exemplified by the E93 gene, which falls at an endpoint in the salivary gland hierarchy because the E93 gene product plays a central role in salivary gland histolysis (Lee et al., 2002). Nevertheless, its expression in other tissues is not responsive to 20E (Baehrecke and Thummel, 1995). The D. melanogaster imaginal discs and ring gland are notable in that they apparently fail to respond to ecdysteroids during the larval/prepupal period, as measured with a GAL4-EcR/GAL4-USP in vivo system in which lacZ staining reports the level of ligand-dependent heterodimerization between the two fusion proteins (Kozlova and Thummel, 2002). The lack of responsiveness may reflect the effect of cofactors (or their absence) and/or mechanisms by which 20E is either removed from the cells via a transporter mechanisms (Hock et al., 2000) or metabolized. The same GAL4 in vivo reporter system indicates that EcR and USP heterodimerization occurs in response to the ecdysteroid peak during D. melanogaster mid-embryogenesis and that the receptor plays a role in germ band retraction and head involution (Kozlova and Thummel, 2003a). Moreover, the transcriptional relationship among ecdysteroidregulated genes is generally maintained during the course of individual ecdysteroid peaks throughout premetamorphic development (Sullivan and

266 The Ecdysteroid Receptor

Thummel, 2003). Mutations of the early-late gene, DHR3, cause embryonic lethality, further implicating not only EcR but the ecdysteroid hierarchy itself (Carney et al., 1997). Of course, the existence of functionally distinct EcR isoforms provides an important basis for differential ecdysteroid responses. During the late larval stage, the EcRA isoform of D. melanogaster is generally predominant in imaginal discs, whereas the B isoforms are associated primarily with larval tissues that undergo histolysis during metamorphosis (Talbot et al., 1993). Issues surrounding the diversity of isoform function will be discussed further in Section 3.5.4.5. 3.5.4.4. Conservation of Ecdysteroid Response Among Insects

The isolation of EcR and USP from other insects and the assembly of their developmental profiles now allows direct analysis of ecdysteroid action in other organisms. So far, these studies show general consistency with those obtained in D. melanogaster, specifically, EcR expression varies over time and peak levels of expression coincide with ecdysteroid peaks, and different isoforms predominate among different tissues. For instance, the expression patterns of the two EcR isoforms in M. sexta vary among cell types (Jindra et al., 1996) and expression levels of the two USP isoforms switch in conjunction with molts (Jindra et al., 1997), providing a regulatory framework for the early-late response of MHR3 discussed earlier (Lan et al., 1999). The recovery of ecdysteroid-inducible target genes such as E74, E75, FTZF1, and HR3 orthologs also allows a comparison of the ecdysteroid-inducible cascade in other organisms, and the resulting proteins play biological roles in a variety of regulatory processes that is too extensive to discuss here. The comparative analysis over preadult development is most complete in M. sexta, and the essential features of the ecdysteroid-inducible hierarchy are maintained (Riddiford et al., 2003, and references therein). Similarly, the players and timing of ecdysteroid-responsive genes during B. mori choriogenesis are conserved (Swevers and Iatrou, 2003), as they are in A. aegypti vitellogenesis (Zhu et al., 2003a) (see Chapter 3.9). Interestingly, however, USP’s other partners, DHR38 and SVP, have not been associated with whole body responses in any organism. In flies, SVP is involved in eye differentiation (Mlodzik et al., 1990) and DHR38 is involved in cuticle formation (Fisk and Thummel, 1998) at the onset of metamorphosis; both players are involved in mosquito vitellogenesis (Zhu et al., 2003a) (see Chapter 3.9).

At least two points emerge from these considerations. First, while the early response at the core of the ecdysteroid hierarchy appears to be highly conserved among insects at the global level, local ecdysteroid responses likely recruit specific players to mediate specific developmental events. Second, combinations of the EcR/SVP/DHR38/USP orphan receptor group might be incorporated into various ecdysteroid-regulated processes for the purpose of regulating a linear series of events such as mosquito vitellogenesis and fly eye differentiation. One intriguing aspect of this orphan group is that the broad spectrum response of DHR38/USP might be important for regulating responses via the ecdysone precursor to 20E and via 20E metabolites occurring after 20E exerts its effects, with SVP providing activation of an alternative pathway by competing with the 20E-inducible response itself. The context of EcR action will be interesting to explore in other insect processes, such as the color pattern formation of butterfly wings (Koch et al., 2003) and insect diapause (see Chapter 3.12). Historically, numerous studies of puffing on polytene chromosomes preceded and have continued along with the work on D. melanogaster. The salivary gland response of Sciara coprophila (fungal fly) provides an interesting comparison to D. melanogaster, and illustrates the importance of investigating the features of each system without pretense. The S. coprophila II/9A puff appearing in response to ecdysteroid treatment reflects the initiation of both bi-directional DNA replication (Liang et al., 1993) and the elevated transcription of two similar mRNA species which bear no resemblance to ecdysteroid-responsive genes found in Drosophila (DiBartolomeis and Gerbi, 1989). When the promoter region of the S. coprophila II/9-1 gene is transgenically introduced into D. melanogaster, it is transcriptionally responsive to ecdysteroids in the salivary gland, indicating that the ecdysteroidinducible transcriptional machinery is conserved evolutionarily between the two species (BienzTadmor et al., 1991). However, there is no evidence that the DNA amplification is induced by 20E in flies or mediated by the ecdysteroid receptor. In Trichosia pubescens, Amabis and Amabis (1984) found that ecdysteroid treatment of salivary glands generates both DNA and RNA ‘‘early’’ puffs, whose subsequent regression is accompanied by the appearance of other ‘‘late’’ DNA amplification and RNA puffs. Cycloheximide treatments after early puff appearance block late puff expansion. In this species, large puffs appear after considerable delay, and the appearance of early puffs is not obvious, suggestive of a single broad ecdysteroid peak, rather

p0455

The Ecdysteroid Receptor

than the two peaks that occur in D. melanogaster over the same developmental timespan. In this respect, the response pattern is comparable to C. tentans, which in turn relies upon an interplay of ecdysone and 20E to achieve ecdysteroid-induced regulation (Stocker and Pavan, 1977). In yet another sciarid fly, Bradysia hygida, 20E induces DNA amplification and transcription of a temporally ‘‘late’’ puff gene (BhC4-1). When this gene is transgenically introduced into D. melanogaster, it retains its ‘‘late’’ puff characteristics in terms of salivary gland timing, but transcription of the gene is not blocked by repression of protein translation with cycloheximide, suggesting that the late induction is actually a form of derepression in the D. melanogaster salivary gland (Basso et al., 2002). This interpretation is also based on the observation that 20E exerts both repressive and inductive effects upon the protein composition of the B. hygida salivary gland during the fourth and final larval stage (de Carvalho et al., 2000). As evidenced by the cases presented here, beyond the superficial similarity, there are numerous unsolved mysteries about the integration of ecdysteroid response into biological processes, especially when one considers that the most complete hierarchical comparisons so far involve only lepidopteran and dipteran species. Several ecdysteroid-inducible genes have been identified for the coleopteran, T. molitor (Mouillet et al., 1999), at least indicating that several of the orphan receptors involved in mediating ecdysteroid responses exist in more primitive insects. Further, the existence of an E78 ortholog in the filarial parasite, D. immitis, implies that at least portions of the hierarchy are conserved in other invertebrates (Crossgrove et al., 2002). 3.5.4.5. In Vivo Molecular and Genetic Analysis of EcR and USP

The pattern of EcR gene transcription during premetamorphic development fluctuates dramatically, with high levels accompanying each of the embryonic and larval molts, followed by peaks of expression during the prepupal and pupal-adult stages of metamorphosis (Koelle et al., 1991). Moreover, the EcR gene encodes three different isoforms with the A isoform’s transcription governed by a promoter and two other isoforms (B1 and B2) generated by alternative splicing via a second promoter (Talbot et al., 1993). All three isoforms share a common sequence that includes a small portion of the A/B domain and the other domains, but the aminoterminal side of the A/B region for each isoform is unique. The A isoform predominates in fly tissues which undergo morphogenetic changes during

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metamorphosis, including the imaginal discs, the ring gland, the imaginal rings of the foregut, hindgut, and salivary gland (Talbot et al., 1993), and in a heterogeneous subset of neurons that degenerate after adult emergence (Truman et al., 1994) (see Chapter 2.4), whereas the B1 isoform is found in larval tissues including the salivary gland, the fat body, larval muscle (Talbot et al., 1993), and proliferative neurons (Truman et al., 1994). Functional studies of EcR based on the isolation of mutations within the D. melanogaster gene have led to further insights about EcR’s developmental role, and essentially confirmed its ecdysteroidmediating function. As might be expected, null mutations of a common region in EcR, and thereby disrupt all three isoforms and result in embryonic lethality, whereas mutations that retain a reduced ability to mediate ecdysteroid responses allow mutant survival through some or all of the larval stages. Some of these weaker mutations affect nonconserved residues, but at least one of these prepupal lethal mutations involves a highly conserved phenylalanine in the DBD; conversely, there are nonconserved residues that cause embryonic lethality when substituted. One of the lethal mutations (A483T) replaces the amino acid residue that interacts with the SMRTER corepressor and is a conditional third instar lethal mutation. Presumably, the lability of the mutant protein disrupts the normal interaction between EcR and SMRTER at higher temperatures (29  C). The A483T mutation (along with other EcR mutations; Table 2) also disrupts adult female fecundity at its restrictive temperature, indicating that the corepressor interaction is essential for at least late larval development and oogenesis (Bender et al., 1997; Carney and Bender, 2000). B-specific mutations cause early larval lethality (Schubiger et al., 1998); an A-specific EcR mutation reduces but does not eliminate mutant survival to the adult stage and disrupts the normal expression of EcRB1 (D’Avino and Thummel, 2000; Schubiger et al., 2003) (see Chapter 2.4). Other A-specific mutations cause pupal lethality (Carney et al., 2004). A gain-offunction mutation of DmEcR (K497A) that dimerizes in the absence of hormone, possibly because of disrupted corepressor binding, has also been identified in cell cultures, though its potential in vivo effect is unknown (Bergman et al., 2004). Over developmental time, the B2 isoform, when expressed under the control of a heat shock promoter, rescues larval development in EcR-null mutants, though heat shocks are required in each instar to accomplish it. The A and B1 isoforms rescue development through the first instar, but fail thereafter, suggesting that a common function is required

268 The Ecdysteroid Receptor

Table 2 A selected list of EcR mutations and their functional consequences. Mutations correspond to shaded residues in Figure 4 Mutation

Species

A393P A398P A522P A534P A559P Y403A A483T

C. fumiferana H. virescens D. melanogaster A. aegypti B. mori H. virescens D. melanogaster

K497A, E Y611F F645A

D. melanogaster D. melanogaster D. melanogaster

W650A

D. melanogaster

Effect

9 > > > > =

Destroys 20E response, but not nonsteroidal agonist response (Kumar et al., 2002; Billas et al., 2003). Impaired coactivator (GRIP1) interaction (Cf EcR only; Kumar et al., 2002)

> > > > ; Reduces nonsteroidal agonist response, but not 20E response (Billas et al., 2003) Conditional late larval lethal mutation; disrupts oogenesis (Li and Bender, 2000); binding site for SMRTER corepressor (Tsai et al., 1999) Constitutive transcriptional activity in cell culture (Bergman et al., 2004) Elevated binding to hsp27 EcRE in presence of ecdysone (Wang et al., 2000) Dimerizes normally but is not 20E inducible; dominant negative lethal mutation in vivo (Cherbas et al., 2003) No dimerization (Cherbas et al., 2003)

during the first instar and more differentiated EcR functions are essential later. The rescued B2 transformants become sluggish during the wandering stage of the late third instar, then immobile, and eventually die (Li and Bender, 2000). Isoform-specific mutations in conjunction with transformation rescue have further delineated EcRbased functions in fly development. As expected, an EcRB1-specific mutation disrupts developmental activities in those cells where it is expressed. For instance, the salivary gland’s ecdysteroid-induced puffing is disrupted in B1 mutants (but not eliminated), and only transformation with EcRB1 restores normal puffing of various early and early-late genes, though B2 exerts a partial rescue. Similarly, B1-expressing abdominal histoblasts and midgut cells develop abnormally (Bender et al., 1997). Neuronal remodeling during metamorphosis is disrupted in genetic mosaics that do not express either of the two B isoforms in proliferating neurons (Schubiger et al., 1998) (see Chapter 2.4), but remodeling is rescued by the expression of either B isoform within these cells (Schubiger et al., 2003). Remodeling is not disrupted in mutations of the early puff genes, indicating that this aspect of the cascade is not specifically tied to the failure of B isoform mutant effects (Lee et al., 2000). A related strategy for discriminating EcR functions involves introducing an isoform or fusion protein transgenically into flies, expressing the transgene ectopically, and then observing the dominant negative phenotypic consequences of such expression. At least three variations have been reported: (1) expression of a wild-type isoform ectopically under the control of an UAS promoter regulated by the yeast GAL4 transcription factor (Schubiger et al., 2003), (2) the expression of a GAL4-EcR LBD fusion protein that forms an

inactive dimer with cellular USP in a nonisoform specific manner (Kozlova and Thummel, 2002, 2003a), and (3) UAS-controlled expression of a dominant negative EcRB1 isoform (F645A in D. melanogaster) that forms an inactive dimer with intracellular USP, and is then tested for rescuability by the concomitant expression of a specific isoform (Cherbas et al., 2003). In the case of wild-type overexpression, each isoform generates a unique pattern of phenotypes. Overexpression of EcRA suppresses posterior puparial tanning, and affects ecdysteroidinducible gene expression in posterior compartments of the wing disc but does not affect viability. By contrast, overexpression of EcRB1 and B2 during puparial formation greatly reduces viability (Schubiger et al., 2003) (see Chapter 2.4). Ectopic expression of a GAL4-EcR during the late third instar causes a failure of puparial contraction and cuticular tanning that resembles the traits displayed by mutant EcR larvae (Kozlova and Thummel, 2002). The isoform-specific rescue of the EcRF645A mutants, however, reveals a poor correlation between tissue-specific effects and intracellular titers of each isoform, though the accumulation of disrupted responses in several individual tissues leads to a stage-specific developmental arrest in the third instar (Cherbas et al., 2003). Further insights about the functional differences associated with the D. melanogaster EcR isoforms will likely emerge by understanding the basis for isoform-specific mRNA transcription. Two promoters, one associated with the A isoform, and the other associated with the two B isoforms, regulate the appearance of these transcripts. Promoter segments responsible for high levels of EcRA expression during metamorphosis have been identified (Sung and Robinow, 2000). While the level of EcRA detected is homogeneous among those neurons which express

The Ecdysteroid Receptor

EcRA during metamorphosis, the underlying promoter regulation is surprisingly heterogeneous among them, indicating that the observed pattern of expression obscures an underlying regulatory complexity. A TGF-b/activin signaling pathway has been implicated in the regulation of EcRB1 transcription. In activin mutants, EcRB1 transcription is reduced and neurons in the mushroom bodies of the brain fail to undergo remodeling during metamorphosis. The expression of EcRB1 rescues the remodeling defects, indicating that EcRB1 levels are regulated through the activin signaling pathway (Zheng et al., 2003). By comparison with EcR’s complexity, the regulation of the usp gene in D. melanogaster is relatively simple, with no introns and no alternative splicing forms, though the profile is more complex in species with multiple USP isoforms (Jindra et al., 1997; Vogtli et al., 1999). In flies, the expression of USP through development is relatively stable, though it is unclear whether EcR or USP is the rate-limiting partner in developing tissues (Henrich et al., 1994). The usp gene is defined by several recessive early larval lethal mutations: three missense substitutions (usp3, usp4, and usp5) that mutate amino acid residues in the USP DBD and directly contact phosphate residues in the DNA backbone, along with a nonsense mutation, usp2, that truncates the DBD (Oro et al., 1990; Henrich et al., 1994; Lee et al., 2000). The null-usp2 allele evokes a different effect on gene expression than the other usp alleles. Whereas all the mutations disrupt the normal repression of the BrC-Z1 isoform, only USP2 is incapable of activating BrC-Z1 transcription. USP3 and USP4 are also able to mediate ecdysteroid-induced gene transcription through an hsp27 EcRE-regulated promoter (Ghbeish et al., 2001). This dual capability has been analyzed in vivo during ommatidial assembly and differentiation in the eye disc. Briefly, the differentiation of the eight retinula cells in each of the 700 or so ommatidia that become the compound eye occurs through the recruitment of undifferentiated cells as a wave moves from the posterior to anterior end of the eye disc. Cells along this progressing wave, the morphogenetic furrow, undergo a host of transcriptional changes and some aspects of the process are ecdysteroiddependent in vitro. Furrow advancement accelerates in mutant usp patches on the eye disc (Zelhof et al., 1997), whereas an ecdysteroid deficit retards its advancement (Brennan et al., 1998). The apparently paradoxical action results from the failure of a repressive usp function as evidenced by the abnormal appearance of Br-C Z1 expression in mutant patches lying along the front of the moving furrow.

269

The usp2 allele causes an absence of Z1 behind the furrow, where it is normally present, further demonstrating the dual roles. USP has been similarly implicated in both wing margin bristle differentation (Ghbeish and McKeown, 2002), and the repression of premature neuronal differentiation in the wing imaginal discs (Schubiger and Truman, 2000) (see Chapter 2.4). The maintenance of USP3 and USP4 activation functions likely explains the normal appearance of mutant usp imaginal clones (Oro et al., 1992). Maternal contribution of normal usp transcript is essential for the completion of embryogenesis (Oro et al., 1992). Mutant usp larvae are rescued through the third instar with a USP connected to a heat shock promoter. The lethal phenotype of these partially rescued larvae is reminiscent of the effects seen with larval EcR mutations and internal morphology is similar at the time of larval arrest. Only the usp mutants, however, develop a supernumerary larval cuticle and fail to wander off the food (Hall and Thummel, 1998; Li and Bender, 2000). DHR38 mutants also undergo abnormal cuticle apolysis (Kozlova et al., 1998), suggesting that the DHR38/ USP dimer may be essential for this aspect of premetamorphic development, rather than the EcR/USP dimer. It is beyond the realm of this chapter to explore the effects of ecdysteroid-inducible genes in depth and the reader is directed to other reviews (Henrich et al., 1999; Riddiford et al., 2000; Thummel, 2002). Isoform-specific mutational analysis of ecdysteroid-responsive genes has generally confirmed essential roles at metamorphosis with the two isoforms of E74 (Fletcher et al., 1995), two of the E75 isoforms (Bialecki et al., 2002), DHR3 (Lam et al., 1999), and the bFTZF1 isoform (Broadus et al., 1999). The complex genetic complementation of several Br-C mutations (Kiss et al., 1988) has been associated with each of four isoforms (Z1–Z4; Bayer et al., 1997). The npr1 (nonpupariation) third instar lethal mutations of Br-C fail to complement all other alleles by impairing a common and essential premetamorphic function. Some Br-C mutations affect the gene’s own expression (Gonzy et al., 2002), consistent with predictions of the Ashburner model. As noted elsewhere, Br-C isoforms play a variety of inductive and repressive roles, often through switchover of isoform expression, in a large spectrum of premetamorphic processes. 3.5.4.6. Ecdysteroid Receptor Cofactors

As noted earlier, the ecdysteroid receptor is part of a complex of proteins which affect both its inductive

270 The Ecdysteroid Receptor

p9025

p9030

and repressive transcriptional functions. In the case of Hsc70, a chaperone that physically interacts with DmEcR, this functional importance is demonstrated by the genetic interaction between EcR and Hsc70 mutations (hsc4 gene) in D. melanogaster. Trans-heterozygotes for these two mutations develop severely blistered wings and bent legs typically associated with the mutational impairment of EcR function (Bender et al., 1997; Arbeitman and Hogness, 2000). Based on findings from experiments with vertebrate nuclear receptors (see Chapter 3.6), transcriptional cofactors and their roles have begun to be recognized and explored; the process of eye ommatidial differentiation during the third instar of D. melanogaster that was described earlier has become the focus of efforts to understand the role of cofactors in development. The dominant negative EcRF645A mutation severely disrupts eye development (Cherbas et al., 2003; Sedkov et al., 2003), and USP function is also required for normal furrow progression (Zelhof et al., 1997). One cofactor implicated in the process is the product of the trithorax-related (trr) gene, a histone methyltransferase that conceivably plays a role in remodeling the chromatin in promoter regions, just as vertebrate factors do to facilitate receptormediated transcription (Stallcup, 2001). The role of TRR as an ecdysone-dependent coactivator is evidenced by the fact that it is recovered as part of a complex that includes EcR and a trimethylated form of the histone-3 protein (presumably modified by TRR) from the ecdysteroid inducible promoter of the Drosophila hedgehog (hh) gene in extracts derived from 20E challenged S2 cells. The methylation is reduced in complexes taken from trr mutant embryos. Further, the trans-heterozygotic combination of a trr mutation and the EcRF645A mutation causes almost complete lethality, revealing an essential in vivo interaction. The aforementioned Hedgehog (Hh) protein regulates the progression of eye cell differentiation, and trr-mutant somatic cell clones express Hh at reduced levels. It follows that TRR is normally a coactivator associated with elevated ecdysteroid-inducible activity leading to higher Hh levels. EcR retains its repressive capabilities in the absence of its inducible activity, and another cofactor, the corepressor SMRTER, has also been implicated in the regulation of Hh expression and eye differentiation. The lethal and mutant phenotypic effects of the dominant negative EcRF645A mutation are reversed by a hypomorphic mutation of the Smr corepressor (i.e., a partial loss of repression), just as EcR mutant effects are overcome by increasing TRR coactivator activity (Sedkov et al., 2003).

Both TRR and SMRTER carry the LXXLL amino acid motifs (L refers to leucine and X refers to any amino acid) associated with nuclear receptor interactions, and physical interaction sites with EcR have been mapped in both cases (Tsai et al., 1999; Sedkov et al., 2003). Moreover, the SMRTER interactive site in DmEcR has been mapped to an amino acid residue in helix 5 that when mutated (A483T) results in conditional larval lethality (Bender et al., 1997; Tsai et al., 1999). Another Drosophila coactivator, Taiman (TAI), was first identified in a screen for mutations that disrupt oogenesis. TAI resembles a human ortholog (AIB1) belonging to the p160 steroid receptor coactivator family (SRC), whose members are typified by a basic helix-loop-helix (bHLH) domain, a PAS domain of unknown function, several LXXLL motifs associated with nuclear receptor interaction, and several glutamine-rich stretches. These proteins form a bridge between hormone receptors, chromatin modifying enzymes such as the histone acetyl transferases, and the transcriptional machinery. TAI expression colocalizes with EcR and USP in the border cells of the ovary and colocalizes with USP on the polytene chromosomes of Drosophila larval salivary glands. TAI also elevates ecdysteroid-inducible transcription in a cell culture and coprecipitates with EcR, but not USP (Bai et al., 2000). Its role in premetamorphic processes has not yet been elucidated. Still another EcR-interacting protein containing the LXXLL motif, rigor mortis (rig), is required for ecdysteroid signaling during larval development. Mutant rig larvae fail to survive beyond the advent of metamorphosis, while displaying phenotypes resembling other mutations defective in ecdysteroid synthesis or response. Rig is required as a coactivator for induction of the E74A isoform which normally appears as ecdysteroid titers increase, but is not required for E75A, EcR, or USP transcription; the effect on transcription is likely to be indirect since Rig contains no DNA-binding motifs. The protein interacts physically with EcR and USP, and also with the orphan receptors DHR3, bFTZ-F1, and SVP. Even when helix 12 is deleted from bFTZ-F1, its interaction with Rig is detectable, suggesting that the relationship between Rig and nuclear receptors is ligand-independent (Gates et al., 2004). A D. melanogaster corepressor, Alien, that is highly conserved phylogenetically, also interacts with several receptors, including EcR, SVP, and bFTZ-F1 (but not DHR3, DHR38, DHR78, or DHR96; Dressel et al., 1999). No mutations of Alien have been reported so far. The pattern of receptor interactions seen with TAI, Rig, and Alien suggests that a level of regulation

p0545

The Ecdysteroid Receptor

remains to be elucidated in connection with the ecdysteroid hierarchy. This is further highlighted by the effects of another cofactor, Bonus (Bon), which belongs to a class of proteins (TIF1) that do not bind to DNA directly, but which repress transcriptional activity. Homozygous bon mutants display many of the defective bFTZ-F1 phenotypes noted earlier (Broadus et al., 1999), and Bon physically interacts with helix 12 of bFTZ-F1 via an LXXLL motif. In some bon mutants, transcript levels of EcR B1, E74A and B, and BR-C are reduced, but DHR3 transcript levels are elevated (Beckstead et al., 2001). Non-DNA binding cofactors, MBF1 and MBF2, associated with Bombyx bFTZ-F1 activity have also been identified which interact with the TATA binding protein to induce transcription (Li et al., 1994). Finally, it is notable that the methopreneresistant (Met) mutation in D. melanogaster defines a gene which specifies another bHLH-PAS transcription factor (Ashok et al., 1998) (see Chapters 3.7 and 3.9). Cellular extracts from flies homozygous for Met mutations show little binding to the juvenile hormone analog methoprene (Shemshidini and Wilson, 1990). The implications of this identity and its significance for explaining the modulatory effects of JH on ecdysteroid-inducible transcriptional activity will be addressed later. 3.5.4.7. Orphan Receptor Interactions with EcR and USP

In addition to those players which have been tied directly to the transcriptional response elicited by the EcR/USP heterodimer (DHR3, E75, E78, bFTZF1), several other nuclear receptors also modulate receptor activity by either dimerizing directly with USP, or by competing for EcR/USP promoter sites. The DHR38/USP interaction is particularly interesting because in vitro results suggest that it is this ‘‘receptor’’ complex that responds to a broad spectrum of ecdysteroids, including many known to exist in D. melanogaster larvae. Further, DHR38 mutations disrupt cuticle formation, as do usp mutations, but EcR mutations do not impair this aspect of development, leaving open the possibility that this aspect of ecdysteroid regulation does not involve EcR at all (Hall and Thummel, 1998; Kozlova et al., 1998). DHR38 is broadly expressed but transcript levels appear to be in low abundance (Sullivan and Thummel, 2003), though the DHR38/ USP heterodimer is more responsive to 20E than EcR/USP (Baker et al., 2003). In any case, the mechanism of action is novel, since the transcriptional

271

activation via DHR38/USP is not associated with a physical ligand interaction. A similar dimer between AHR38 and AaUSP has been noted prior to vitellogenesis in A. aegypti. The nonresponsive heterodimer is displaced by the EcR/USP heterodimer in response to a 20E titer peak following a bloodmeal (Zhu et al., 2000) (see Chapter 3.9). As noted earlier, USP also dimerizes with SevenUp (SVP), the aforementioned ortholog of COUP-TF. As the 20E peak that stimulates A. aegypti vitellogenin expression ensues, a similar competition between EcR and SVP for USP as a heterodimeric partner leads to a downregulation of A. aegypti vitellogenin gene transcription after egg-laying (Zhu et al., 2003a), illustrating a mechanism by which an ecdysteroid response can be terminated. Ectopic expression of the svpþ gene in flies causes lethality, but this effect can be rescued by simultaneous ectopic expression of uspþ (Zelhof et al., 1995a). This interaction apparently is a relevant aspect of photoreceptor differentiation, since both SVP and USP function are essential for this process to occur normally (Mlodzik et al., 1990; Zelhof et al., 1997). The potential relevance of the SVP/USP interaction has also been established by demonstrating that SVP competes with EcR for USP’s partnership, and thereby reduces ecdysteroid-induced transcription in cell cultures. Further, the functional SVP/USP dimer preferentially interacts with DR1 DNA elements, whereas the EcR/USP dimer interacts with the Eip28/29 element. Still another orphan receptor that is essential for normal metamorphosis in flies is DHR78. Mutations of this receptor cause late larval lethality, disruptions of the tracheal system, and the impairment of EcR, E74B, and BR-C transcription. DHR78 binds to numerous ecdysteroid inducible sites in salivary glands (Fisk and Thummel, 1998; Astle et al., 2003). DHR78 inhibits 20E-dependent induction of transcription in cell culture assays (Zelhof et al., 1995b) and it has been proposed that DHR78 activity may interact with an ‘‘upstream’’ ligand in the period that precedes the late third instar ecdysteroid peak to prime a later ecdysteroid response (Fisk and Thummel, 1998). A Bombyx ortholog, BHR78, has been shown to dimerize directly with BmUSP (Hirai et al., 2002) suggesting that DHR78 may block 20E-dependent transcription by competing for USP with EcR. 3.5.4.8. Ecdysteroid Action and Other Developmental Processes

The ecdysteroid response is highly heterogeneous among cell types, owing to differences in the

272 The Ecdysteroid Receptor

quantitative levels of EcR and USP isoforms, rates of 20E conversion from ecdysone, 20E metabolism and cellular exclusion, and yet it is evident that the circulation of ecdysteroids in the insect hemolymph provides the organism with a means to coordinate its individual developmental programs. Therefore, it is expected that ecdysteroids set off general responses and also specific responses that involve either EcR and USP or targets that are regulated by them. For organismal investigations, the dramatic changes associated with the late larval ecdysteroid peak in D. melanogaster evoke substantial changes in transcript levels which can be detected using genomic approaches. As already noted, a combination of standard technology and a genomic outlook has motivated the assembly of a detailed profile of orphan receptor transcript levels through preadult development (Sullivan and Thummel, 2003). A limitation of the whole body approach is evident in the case of DHR38, which is widely expressed but at such low levels that the transcript is barely evident on Northern blots. An earlier study using subtractive hybridization led to the characterization of several ecdysteroid-inducible genes, most of which were unknown and not associated with ecdysteroidinducible events directly (Hurban and Thummel, 1993). Screens of hematopoietic Kc cells (Savakis et al., 1980) and imaginal discs (Apple and Fristrom, 1991) have yielded several ecdysteroid-dependent genes which do not overlap with those recovered from other screens, and these transcripts are largely associated with tissue-specific responses. However, while the timing of early ecdysteroid gene expression is delayed in the discs (Huet et al., 1993), at least one ecdysteroid-responsive pupal cuticle gene, EDG84A, is regulated by FTZ-F1, which as noted, also regulates metamorphic processes in the salivary gland (Murata et al., 1996). Microarray analysis has been used more recently to investigate the changes in transcription on a genome-wide basis, using the onset of the white prepupal (puparial) stage as a reference point. The early puff genes change in a predictable, ecdysteroid-regulated manner, and genes involved in processes such as myogenesis, apoptosis, and imaginal disc differentiation are upregulated at appropriate times during the period. As ecdysteroid levels peak, transcription of most genes encoding glycolytic enzymes are substantially reduced, as are genes whose enzymatic products regulate the citric acid and fatty acid cycles, oxidative phosphorylation, and amino acid metabolism (White et al., 1999). Adult flies heterozygous for a lethal EcR mutation survive longer than wild-type flies and show greater resistance to oxidative stress, heat, and dry

conditions, though activity levels are normal and a possible connection to metabolic function has not been made (Simon et al., 2003). Important mechanistic activities at the cellular level, however, will continue to require careful functional analysis. In the case of eye differentiation in flies, morphogenetic furrow movement during the late third instar is dependent on 20E, but not on normal EcR gene activity (Brennan et al., 2001). The repressive action of USP described earlier for this process (Ghbeish and McKeown, 2002) is offset by the positive regulation of Hedgehog (Hh) transcription that involves EcR (Sedkov et al., 2003). Hh, in turn, facilitates a switchover from the Z2 to the Z1 isoform at the ecdysteroid-responsive BR-C gene locus. The temporal coordination of hormonal signaling with the regulatory activities along the furrow, therefore, provide a balance of signals that both stimulates morphogenetic furrow movement (in the form of EcR-mediated Hh expression) and regulates its rate of progression (in the form of a repressive USP function). The mechanism of furrow movement is not fully elucidated, and it is conceivable that a second hormonal signaling pathway is involved in the process. The interaction between JH and ecdysteroid action remains elusive, despite the clear recognition that there must be cross-talk between these pathways. The JH analog methoprene induces several disruptions of the central nervous system that resemble those caused by mutations of the BR-C gene. These defects are not seen in flies homozygous for the methoprene-resistant (MET) mutation (Restifo and Wilson, 1998). 3.5.4.9. Juvenile Hormone Effects upon Ecdysteroid Action

A central tenet of insect biology is that ecdysteroids mediate larval-larval molts in the presence of the sesquiterpenoid, JH (see Chapter 3.7). In the absence of JH, by contrast, ecdysteroid induces a larval-pupal transition. Implicit in this view is the recognition that JH via its receptor competes with the EcR/USP dimer at an EcRE (Berger et al., 1992) or modifies components of the ecdysteroid receptor itself (Jones et al., 2001; Henrich et al., 2003). Nevertheless, no uniform explanation for the action of JH, alone or in conjunction with ecdysteroids, has been forthcoming. The structural similarity of 9-cis retinoic acid, the cognate ligand of vertebrate RXR, with JH inspired experiments to test the possibility that the insect RXR ortholog, USP, is the JH receptor, and like RXR, switches between a homodimeric and heterodimeric state to process incoming hormonal

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signals. This enticing possibility is further bolstered by the observation that RXR is inducible by binding to the acid metabolite of the JH analog, methoprene (Harmon et al., 1995). Jones and Sharp (1997) reported that a Drosophila USP fusion protein binds to JH forms found in flies, as measured by the dose-dependent suppression by JH3 and JH3-bis epoxide on the fluorescence of tryptophan residues in the ligand binding domain of USP. Later studies have demonstrated that purified USP fusion proteins exist in homodimeric and homotetrameric forms which show a saturable ability to bind to JH3, with about 50% binding at 4 mM, while the closely related farnesol exerts no effect. Using nuclear extracts from the lepidopteran Sf9 cell line, it was further shown that USP binds specifically as a homodimer to a direct repeat response element separated by 12 nucleotides (DR12). Further, JH3 strongly induces a reporter gene in which the DR12 element is attached to the core promoter of the gene encoding the Trichoplisia ni JH esterase (Jones et al., 2001). A DR4 element in the C. fumiferana JH esterase gene promoter is induced by JH1 and suppressed by 20E, suggesting that an interaction between these hormones occurs at the DNA level (Kethidi et al., 2004). The biochemical examination of these promoters, along with those of JH-inducible genes found recently in cell cultures through microarray analysis, will hopefully bring a convergence of views on JH action (Dubrovsky et al., 2000). Along a different line of reasoning, experiments have been undertaken based on the sequence similarity between the EcR LBD and the vertebrate FXR LBD, which surprisingly, is responsive to JH3 in the 10–50 mM range with an RXR dimer partner in mammalian cell cultures (Forman et al., 1995). When tested in this regime, JH3 potentiates the ability of the EcR/USP complex to induce a weak transcriptional induction via an hsp27 EcRE in the presence of submaximal muristerone A levels. The potentiation apparently requires prior binding of EcR to ecdysteroids and there is no evidence yet that JH3 is binding with USP in this assay, and no response is noted with EcR/USP to bile acid, the most potent FXR activator. When tested with 20E, only EcRB2 among the three Drosophila isoforms is additionally potentiated with JH3 (Henrich et al., 2003), and, as noted, the B2 isoform is the only one capable of rescuing larval development in EcR mutants (Li and Bender, 2000). The potentiation is not analogous to the responsiveness of an EcR/RXR heterodimer described earlier (Saez et al., 2000), and the low ecdysteroid levels which accompany JH titers during larval development suggests its possible biological relevance. In both Drosophila

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and Manduca, one of the isoforms of Broad (Z1) is ectopically induced in cuticle by the application of JH analogs, further supporting the possibility that JH acts directly upon the ecdysteroid receptor’s transcriptional capability (Zhou and Riddiford, 2002), since the BR-C gene is a direct target of the receptor in both Drosophila and Manduca. In reality, differences such as the promoter and receptor constructs among the experimental regimes leave open the possibility that USP acts by multiple mechanisms, particularly if one considers the possibility that posttranslational modifications such as phosphorylation also can modify receptor activity. Possibly distinct roles of USP during larval and metamorphic development are evidenced in vivo by the fact that a chimeric USP transgene (in which the Drosophila USP LBD is replaced by the equivalent domain from C. tentans) completely rescues larval development in usp mutants that otherwise die in the first instar. Such genetically rescued larvae, however, suddenly die as the onset of the prepupal stage approaches (Henrich et al., 2000). Another potentially important factor is the protein defined by the Methoprene-tolerant (Met) mutation that belongs to the bHLH-PAS family of transcription factors, and includes several cofactors known to interact with nuclear receptors (Ashok and Wilson, 1998). The effects of Met upon the ecdysteroid receptor await further experimentation in Drosophila and other insects. JH3 plays an essential role for the translation of bFTZ-F1 from existing transcripts in the fat body of newly emerged A. aegypti females. In turn, FTZF1 activity is required for the subsequent transcription of the mosquito ‘‘early puff’’ genes regulating vitellogenesis (Zhu et al., 2003b) (see Chapter 3.9). While this may be seen as another mode of action, the effect on bFTZF1 translation may be indirect, that is, JH3 may regulate the transcription of genes whose products play a role in mRNA stability and/ or protein translation. In summary, the diverse functional effects of JH upon ecdysteroid-inducible gene expression have elicited several possible mechanisms from investigators over the years. As it becomes increasingly apparent that EcR and USP are the two most recognizable components of a protein complex of diverse functional capability, it seems highly plausible that there is not a single JH receptor, but rather, a JH receptive protein complex (or complexes).

3.5.5. Prognosis An explosion of information about the ecdysteroid receptor has occurred recently which will

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undoubtedly influence the field far into the future. There are three levels of progress that will be central to the development of the field: (1) continued progress on understanding the biological action of ecdysteroids using functional genomics and genetics in D.melanogaster, (2) the adaptation of transgenic and genomic methods into receptor studies of other insects, and (3) continued modeling of insect receptor complexes along with tests of these models through both in vitro and in vivo experimentation. The emergence of clever new methodologies, notably the in vivo lacZ reporter system reported by Kozlova and Thummel (2003b), provides in vivo tools that overcome the limitations of biochemical isolation and testing from flies. Microarray analysis has already been employed to investigate changes in gene transcription at the onset of metamorphosis (White et al., 1999) and to compare expression in Kc167 and SL2 cell lines (Neal et al., 2003). Microarray analysis has also been employed in D. melanogaster to follow expression events throughout larval and metamorphic development. The results suggest that while the larval stage is relatively quiescent, there are unique patterns of expression associated with this early developmental time which in some cases, may prove to be repressive functions (Arbeitman et al., 2002). In this respect, microarrays also may prove important for gaining a better grasp of JH’s mode of action (Dubrovsky et al., 2000) during larval stages. Inevitably, the genomic approach will unravel entire gene networks that interact with and operate within the ecdysteroidregulated hierarchy (Arbeitman et al., 2002; Stathopoulos and Levine, 2002). In any event, investigating the entire genome is important for identifying temporal connections at a gene expression level which potentially coincide with important functional relationships, as the connection between the glycolytic pathway and the onset of metamorphosis has already exemplified (White et al., 1999). Sooner rather than later, it is likely that genomewide approaches will be applied to identify EcR and/or USP binding sites (Ren et al., 2000). Another feature of ecdysteroid action which has received virtually no attention is the role that nutritional and cellular states play in determining and modifying ecdysteroid action (Zinke et al., 2002). As the case with glycolysis shows, it seems inevitable that a relationship exists here, given that the most important problem for premetamorphic insects stems from their need to process nutrients as they grow and undergo molting. The role of orphan receptors and their ligands seems an obvious place to explore this possibility, since most of the identified orphans among vertebrates have proven to be

responsive to a variety of dietary compounds and intracellular metabolites, including EcR’s vertebrate relatives, FXR and LXR. The regulation of EcR and USP expression has begun to be undertaken and should also lead to important insights about endocrine regulation generally, and the interaction of ecdysteroid regulation with other biological processes (Sung and Robinow, 2000; Zheng et al., 2003). EcR undergoes fairly dramatic fluctuations in transcription, though little is known about the relative titers of EcR and USP intracellularly (Koelle et al., 1991). In other words, it is not clear which one of the two is rate-limiting in any given cell type, and thus, a quantitative question remains to be fully considered. In fact, there has been a tendency to ascribe regulation to the roles of individual players, without regard to the fact that cellular protein titers, ligand titers, rate of metabolism, and the relative affinity of promoter elements all play a role in determining the level of transcriptional response at any given gene at any given time. When ecdysteroid action is viewed as a challenge of cellular equilibrium, it is seen as a continuous and interactive process rather than a directed and responsive one. Interestingly, there are still unanswered questions about ecdysteroid regulation within the Drosophila salivary gland hierarchy which will be particularly useful for examining ecdysteroid response and processing, particularly with the judicious use of transgenic constructs. A second important cornerstone for future progress will depend upon a more complete depiction of ecdysteroid receptor function among the insect orders. A survey of known EcR and USP sequences shows that only Diptera and Lepidoptera are extensively represented at this time. Based upon the variations in agonist responsiveness already discovered among the insect receptors in these two orders alone, the need for obtaining and testing other insect receptors is plainly obvious. The ability to assess the effects of EcR, USP, and genes targeted by the ecdysteroid receptor in other insects will be greatly enhanced by continued development of transformation procedures. The ability to produce ‘‘null’’ mutations via RNA interference with transgenic constructs will be particularly important for assessing functional processes in other insects. Uhlirova et al. (2003) illustrated this possibility by showing that Sindbis virus-induced transformation with an RNAi eliminates Br-C expression in Bombyx mori. The interference exacerbates the same developmental defects in Bombyx as previously noted for the effects of Br-C null mutations in flies. Related to the discovery and characterization of ecdysteroid responses in insect processes will be the

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continued examination of the insect receptors themselves. The use of chimeras and mutational analysis, in conjunction with the predictive powers provided by improved modeling and crystal structures, will lead to further insights concerning the capabilities of receptor function and its response to ligands, which may include a broad spectrum based on the flexibility of its ligand-binding pocket (Billas et al., 2003). Many mutations affect receptor function nonspecifically (Bender et al., 1997; Bergman et al., 2004), and site-directed mutagenesis provides a particularly useful approach for finding mutations that specifically disrupt EcR and USP subfunctions essential for interpreting receptor function in cellular and in vivo systems. In summary, the information reported here represents only a fraction of the progress on ecdysteroid receptor action that has taken place over the last 20 years. The convergence of research interests mentioned at the outset has created a synergy among approaches that will continue to yield important new insights about endocrine action and its effects on development, new possibilities for insecticidal discovery, and new applications for the use of ecdysteroid-regulated transcriptional cell systems.

Acknowledgments The author wishes to thank Drs. Dino Moras, Markus Lezzi, and Fraydoon Rastinejad for contributing illustration figures for this chapter. The author also acknowledges Drs Margarithe SpindlerBarth, Klaus Spindler, Markus Lezzi and JeanAntoine Lepesant for useful discussions related to the material presented in this manuscript, along with Dr. Larry Gilbert for his advice on the writing and his friendship. The author is responsible for any inaccuracies in the content and apologizes for the exclusion of many excellent contributions because of space constraints. VCH is currently supported by the US Department of Agriculture’s Competitive Grants program (2003-35302-13474).

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Drosophila RXR, in morphogenetic furrow movement and photoreceptor cluster formation. Development 124, 2499–2505. Zelhof, A.C., Yao, T.P., Chen, J.D., Evans, R.M., McKeown, M., 1995a. Seven-up inhibits ultraspiraclebased signaling pathways in vitro and in vivo. Mol. Cell. Biol. 15, 6736–6745. Zelhof, A.C., Yao, T., Evans, R.M., McKeown, M., 1995b. Identification and characterization of a Drosophila nuclear receptor with the ability to inhibit the ecdysone response. Proc. Natl Acad. Sci. USA 92, 10477–10481. Zheng, X., Wang, J., Haerry, T.E., Wu, A.Y-H., Martin, J., et al., 2003. TGF-B signaling activates steroid hormone receptor expression during neuronal remodeling in the Drosophila brain. Cell 112, 303–315. Zhou, Z., Riddiford, L., 2002. Broad specifies pupal development and mediates the ‘status quo’ action of juvenile hormone on the pupal-adult transformation in Drosophila and Manduca. Development 129, 2259–2269. Zhu, J., Miura, K., Chen, L., Raikhel, A.S., 2000. AHR38, a homolog of NGF1-B, inhibits formation of the functional ecdysteroid receptor in the mosquito Aedes aegypti. EMBO J. 19, 253–262. Zhu, J., Miura, K., Chen, L., Raikhel, A.S., 2003a. Cyclicity of mosquito vitellogenic ecdysteroid-mediated signaling is modulated by alternative dimerization of the RXR homologue, Ultraspiracle. Proc. Natl Acad. Sci. USA 100, 544–549. Zhu, J., Chen, L., Raikhel, A.S., 2003b. Posttranscriptional control of the competence factor bFTZ-F1 by juvenile hormone in the mosquito Aedes aegypti. Proc. Natl Acad. Sci. USA 100, 13338–13343. Zinke, I., Schutz, C.S., Katzenberger, J.D., Bauer, M., Pankratz, M.J., 2002. Nutrient control of gene expression in Drosophila: microarray analysis of starvation and sugar-dependent response. EMBO J. 21, 6162–6173.

3.6 Evolution of Nuclear Hormone Receptors in Insects V Laudet and F Bonneton, Ecole Normale Supe´rieure de Lyon, Lyon, France ß 2005, Elsevier BV. All Rights Reserved.

3.6.1. Introduction 3.6.2. Nuclear Receptors in the Animal Kingdom 3.6.2.1. The Phylogeny of Nuclear Receptors 3.6.2.2. Metazoan Phylogeny and Nuclear Receptors: the Ecdysozoan Problem 3.6.3. Phylogeny of Nuclear Receptors in Arthropods 3.6.3.1. Towards an Inventory 3.6.3.2. Conservations and Innovations 3.6.4. Evolution of the Ecdysone (20E) Receptor 3.6.4.1. The Arthropod Ecdysone Receptor is a Heterodimer between ECR and USP-RXR 3.6.4.2. Divergence of the Ecdysone Receptor in Diptera and Lepidoptera 3.6.4.3. Evolution of USP-RXR 3.6.4.4. Evolution of ECR 3.6.4.5. Ecdysone Receptor and the Evolution of Insects

3.6.1. Introduction p0005

Nuclear receptors form a superfamily of ligand activated transcription factors, which regulate various physiological functions from development or reproduction to homeostasis and metabolism (Laudet and Gronemeyer, 2002). This superfamily is present in all metazoans, and only in metazoans – no nuclear receptors have been found in the complete genome sequences currently available for plants, fungi, or unicellular eukaryotes (Escriva et al., in press). In insects, nuclear receptors are implicated in embryogenesis as well as in the control of the postembryonic development, namely molting and metamorphosis (Kozlova and Thummel, 2000) (see Chapters 3.3 and 3.5). One of the most important insect hormones, 20-hydroxyecdysone (20E) is a ligand for a nuclear receptor, the ecdysone receptor (EcR) (Koelle et al., 1991) (see Chapter 3.5). Although EcR is commonly called the ecdysone receptor, it is really the receptor for the principal molting hormone of insects, 20E. Ecdysone is in most cases the precursor of 20E. Although it is strongly debated, it is interesting to note that several studies have suggested that juvenile hormone (JH), a hormone that counteracts the action of ecdysone, might also act through nuclear receptors (Jones and Sharp, 1997) (see Chapter 3.7). It is of note that the superfamily contains not only receptors for known ligands but also a large number of so-called orphan receptors for which ligands do not exist or have not been identified. It

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is not known whether all of these orphan receptors indeed have a ligand, if they activate transcription in a constitutive manner, or if they have alternative transcriptional regulation mechanisms (Giguere, 1999). In insects, many orphan receptors are direct target genes of the EcR and participate in the gene regulatory hierarchy that controls metamorphosis. Nuclear receptors are modular proteins, with three major domains arranged linearly from the N-terminus to the C-terminus (Figure 1). At the N-terminus there is a transactivation domain, most often called AF-1, which is of variable length and sequence in the different family members and which is recognized by other proteins in the transcription complex. The DNA binding domain (DBD), the site for specific interaction with DNA, is in the mid region of the protein and harbors two zinc finger motifs common to all members of the family. This domain is absent in two divergent members of the superfamily DAX-1 and SHP that are found only in vertebrates (Laudet and Gronemeyer, 2002). The C-terminal contains the ligand binding domain (LBD), whose overall architecture is well conserved between the various family members but which is nonetheless different enough to permit the high degree of ligand specificity characteristic of each member of the family. This domain harbors a ligand dependent activation function, called AF-2, that is responsible for the recruitment of transcriptional coactivators. As with DBD, the LBD is absent in

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Figure 1 Schematic illustration of the structural and functional organization of nuclear receptors. The evolutionary conserved regions C and E are indicated as boxes and a black bar represents the divergent regions A/B, D, and F. Note that region F may be absent in some receptors. The functions of the domains are depicted below and above the scheme; most of these are derived from analyses of vertebrate receptors. Within the transcription activation functions, AF-1 and AF-2, autonomous transactivation domains have been defined in most receptors. These are depicted as circles in the A/B and E domain. The activation domain of the E region has been mapped within the C-terminal helix 12. Note that there are many receptor genes that encode different isoforms harboring different A/B regions, and thus different AF-1 and activation domains.

some divergent members of the superfamily such as the Knirps group in insects (Laudet and Gronemeyer, 2002). The three-dimensional structure of the LBD has been determined for several nuclear receptors, including both unliganded (apo) or liganded (holo) forms, allowing increased understanding of the mechanisms involved in ligand binding and transactivation functions (Figure 2). Interestingly, the structure of the EcR-USP LBD heterodimer from the moth Heliothis virescens has recently been established (Billas et al., 2003). These crystal structures show that the E domain undergoes a major conformational change upon ligand binding, allowing the interaction with coactivators and the transactivation of target genes. Many nuclear receptors are transcriptional silencers in the absence of ligand (apo-receptor) as a result of interaction with intermediary factors (i.e., corepressors). The determination of the crystal structure of nuclear receptors has shown that the LBD region is made up of a threelayered a-helical antiparallel sandwich of 11–12 helices forming a hydrophobic pocket (Wurtz et al., 1996). Upon ligand binding (holo-receptor), the ligand makes different contacts with amino acid residues, promoting a conformational change that closes the ‘‘lid’’ (helix 12, H12) on the pocket and the corepressor complex dissociates. Thus, the activation domain within the H12 (AF2-AD) is able to interact with coactivators and promotes transcription of target genes. The conformational change of the LBD upon ligand binding is therefore necessary for the transactivation function of nuclear receptors

(Steinmetz et al., 2001; Laudet and Gronemeyer, 2002). Nuclear receptors bind as homodimers or heterodimers to the regulatory regions of target genes (usually to the sequence PuGGTCA) called the hormone response element (HRE) (Laudet and Gronemeyer, 2002) (see Chapter 3.5). However, mutation, extension, and duplication, as well as distinct relative orientations of repeats of this motif, generate response elements that are selective for a given (class of) receptor(s). For example, steroid receptors bind as homodimers to HREs containing two core motifs separated by 1–3 nucleotides and organized as palindromes. Other receptors (e.g., RARs, TRs, HR38) heterodimerize with retinoid X receptors (RXRs, homologous to the insect USP gene) and bind to direct repeats (DRs) of the core motif. The spacing between the two halves of the DR dictates the type of heterodimer that will bind (e.g., a DR separated by five nucleotides (DRs) will be recognized by RXR:RAR and a DR4 by RXR:TR). Some receptors, such as the FTZ-F1, HR3, or E75 orphan receptors, can bind to DNA as monomers through a single core sequence. In this case, an A/T-rich region 50 to the core element governs the binding specificity (Laudet and Adelmant, 1995). The development of nuclear receptor research has largely been driven using data from mouse and Drosophila and it is interesting to note that the knowledge from these two organisms are largely complementary. In the mouse, many in vitro functional data are available and much effort has been deployed to analyze the mode of action of nuclear

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Figure 2 Three-dimensional structure of RXR in the apo or holo form. There are 11 helices that form a compact structure with a ligand binding pocket, the size of which varies between the various family members. The entrance to the pocket is guarded by a twelfth helix (H12), which forms a movable lid over the pocket and contains residues critically required for the function of AF2. The orientation of H12 is determined by the size and shape of the ligand occupying the pocket. In most cases, without ligand the LBD interacts with corepressors through a region called the CoRNR box in the region of helices H3 and H4. The ligand promotes a complex conformational change inducing the repositioning of H12, modifying the CoRNR box and thus leading to corepressor complex dissociation. This repositioning of H12 also allows the recruitment of coactivators involving the interaction with short LxxLL-like motifs called ‘‘NR boxes’’ present in most coactivators. The LxxLL motif recognizes a hydrophobic cleft formed by H3, H4, and H12 that has an important role in stabilizing this LxxLL-LBD interaction. The conformational change of the LBD upon ligand binding is therefore necessary for the transactivation function of NRs. (Data from Bourguet, W., Ruff, M., Chambon, P., Gronemeyer, H., Moras, D., 1995. Crystal structure of the ligand-binding domain of the human nuclear receptor RXR-alpha. Nature 375, 377–382; and Egea, P.F., Mitschler, A., Rochel, N., Ruff, M., Chambon, P., et al., 2000. Crystal structure of the human RXRa ligand-binding domain bound to its natural ligand: 9-cis retinoic acid. EMBO J. 19, 2592–2601.)

receptors. Indeed, much of our knowledge on how nuclear receptors modulate transcription comes from studies on mammalian cell systems. In contrast, Drosophila work has provided essentially an in vivo oriented picture of the developmental role of nuclear receptors. Nevertheless, two important perspectives are missing from the field. First, very few structure/function analyses are available for insect (including Drosophila) nuclear receptors, most often what we know of their mode of action being derived from data obtained on mammals. This obviously hampers our global understanding of insect nuclear receptor action in vivo. Second, most of the data concerning nuclear receptors in insects arise from very few model systems, mainly Drosophila and other Diptera or lepidopterans such as Bombyx mori and Manduca sexta. Taking these limitations into account, this chapter aims to put our current knowledge of insect nuclear receptors in a broader evolutionary perspective. This will be done first by discussing insect nuclear receptors in the context of metazoan phylogeny and evolution of the whole nuclear receptor superfamily.

In the second part, we focus on the evolution of nuclear receptors in arthropods, providing specific data on liganded and orphan receptors in this phylum. Nevertheless, we must emphasize that an enormous amount of functional, genetic, and structural data is available on nuclear receptors and it is unrealistic to attempt to present a complete view of this exponentially expanding field. The interested reader can find several excellent reviews (Truman and Riddiford, 2002; Thummel, 2001 among others, as well as many chapters in this volume (see Chapters 3.7 and 3.9)) and the web specific information in Nurebase, a database specializing in nuclear receptors maintained in Lyon (Nurebase; Duarte et al., 2002; Ruau et al., 2004).

3.6.2. Nuclear Receptors in the Animal Kingdom 3.6.2.1. The Phylogeny of Nuclear Receptors

3.6.2.1.1. Classification of the nuclear receptors Molecular phylogeny has been applied to better delineate the evolutionary history of the nuclear

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receptor superfamily (Amero et al., 1992; Laudet et al., 1992; Laudet, 1997; Thornton and DeSalle 2000; review: Escriva Garcia et al., 2003). Since two main functional domains, the DBD and the LBD, are conserved in most nuclear receptors, phylogenetic trees have been produced using each of these domains alone or in combination. It is now clear that the DBD alone is too short and too conserved to contain a significant phylogenetic signal. Thus, if DBD-based trees contain accurate information on the grouping of closely related receptors (such as TRs and RARs) they are unable to correctly decipher deep relationships. Using the LBD alone provides a better resolution of the phylogeny but this is less robust than the phylogenies arising from the use of both DBD and LBD. Thus, despite several attempts it has been very difficult to compare the evolution of the DBD and the LBD using their phylogenies. The question of some degree of independence between the two domains during evolution is still open (see conflicting views in Escriva et al., 2000 and Thornton and DeSalle, 2000). All the trees discussed in this review were constructed using both domains. It is important to note that several phylogenetic methods, namely maximum parsimony, distance analysis using neighbor-joining, or probabilistic approaches using maximum likelihood, have been used to produce nuclear receptor phylogeny. Despite some differences in specific regions of the trees (respective placement of ER, ERR, and steroid receptors, for example; Laudet, 1997; Thornton et al., 2003), the topology of these trees and the conclusions that can be drawn from them are very similar. Thus, the phylogeny of the nuclear receptor superfamily is now relatively consensual and widely accepted. Based on these results, the family has been divided into six subfamilies that are supported by high bootstrap values (Figure 3). It is worth noting that the relationships between these subfamilies are not supported and it is impossible to say which one is the most ancient, based only on sequence phylogeny. 1. The first subfamily contains several important liganded groups of nuclear receptors such as the thyroid hormone receptors (TRs), the retinoic acid receptors (RARs), the PPARs, the xenobiotic receptors PXR and CAR with their homolog in insect DHR96 (which is still considered an orphan receptor), the bile acid receptor FXR, the ecdysone receptor (ECR), and the oxysterol receptors (LXRs). In addition, this subfamily also contains two groups of orphan receptors, Rev-erb/E75 and ROR/HR3. It is interesting to note that ligands (namely cholesterol and retinoic

2.

3.

4.

5.

6.

acid) were recently proposed for the RAR-related orphan receptor (ROR) (Stehlin-Gaon et al., 2003; Kallen et al., 2002). The relationship between these groups is poorly resolved: if TR and RAR appear very often to cluster together with weak bootstrap values, such as for REVERB and ROR, the only clear and robust grouping joins LXR, FXR, and PXR groups. The second subfamily clusters RXR, the 9-cis retinoic acid receptor, and its arthropod homolog USP that apparently does not bind this compound, and many groups of orphan receptors: TR2 and TR4 and their Drosophila homolog HR78, HNF4, the TLL group (with four genes in Drosophila: TLL, DSF, PNR, and FAX1), and the COUP-TF/SVP group. Of note is the presence of unusual receptors that contain only the LBD sequence, DAX and SHP clearly being members of this subfamily, probably related to TLL receptors (Laudet, 1997). The third subfamily clusters steroid receptors (SRs), the estrogen receptor (ER), and the estrogen related receptors (ERRs) that are still orphan receptors. There is one ERR homolog in Drosophila but not for SRs and ERs. As their names indicate ERRs appear more closely related to ERs than to steroid receptors but this view has been challenged recently by data suggesting that ERs and SRs cluster together and are then joined by ERR. The respective placement of SRs, ERs, and ERRs is an important issue regarding ligand binding evolution and is still under discussion. The fourth subfamily contains only the NGFIB group of orphan receptors with their unique arthropod homolog HR38. The fifth subfamily contains the orphan receptor SF1 group, which contains a Drosophila homolog, FTZ-F1, as well as another gene, DHR39, whose origin is still unclear. The last subfamily contains the orphan receptor GCNF with its ortholog in Drosophila, HR4.

It is important to note that the placement of three Drosophila genes that contain no LBD (KNI, KNRL, and EAGLE) is still unclear. They may be related to HR96 but, given the poor phylogenetic information present in the DBD sequences, it will be difficult to resolve their origin using phylogenetic analysis alone (Laudet, 1997). Given the plethora of names available for the same sequence, it has proved useful to construct a nomenclature based on the nuclear receptor sequences using molecular phylogeny (Nuclear Receptor Nomenclature Committee, 1999). The system is based on the nomenclature system that was developed

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Figure 3 Phylogenetic tree of human (black) and Drosophila (red) nuclear receptors. In each case the chemical nature of the endogenous ligand has been indicated on the right. Orphan receptors are without any indication. This phylogeny shows that there is no relationship between the ligand binding ability of NRs and their evolutionary history. (Reproduced with permission from Robinson-Rechavi, unpublished data.)

for cytochrome P450. Each receptor is described by the letters NR (for ‘‘nuclear receptor’’) and a threedigit nomenclature: the subfamily to which a given receptor belongs is indicated by Arabic numbers,

groups by capital letters, and individual genes by Arabic numbers. Thus, the Drosophila EcR that belongs to subfamily I, to group H in this subfamily, and which is the first gene described in that group

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will be called NR1H1. The receptors that contain only one of the two functional domains (KNI group, DAX/SHP, and some nematode receptors such as Odr-7) are artificially clustered in subfamily 0. This system has proven to be flexible enough to integrate nuclear receptors from invertebrates as well as sequences generated from genome projects for which no pharmacological data are yet available. In order to follow the evolution of this nomenclature, the new names are available on a specific website (http://www.ens-lyon.fr/LBMC/laudet/nomenc. html) as well as on Nurebase. 3.6.2.1.2. Evolution of nuclear receptor functions It is always interesting to compare results from phylogenetic analysis with functional or morphological features of the group of sequences or organisms that are compared. For nuclear receptors, evolution of the superfamily with major functional characterizations of the receptors, namely DNA binding, dimerization abilities, and ligand binding (reviews: Escriva et al., 2000; Escriva et al., in press), is compared. As discussed above, the vast majority of nuclear receptors recognize the core sequence RGGTCA organized either as DRs, inverted repeats (mostly palindromes), or alone with an upstream A/T-rich sequence. Nuclear receptors bind to these elements as homodimers, heterodimers, or monomers. In most cases, heterodimers are formed with RXRs (or USP in insects). When these functional characteristics are compared with the tree there is a very good correlation with the evolutionary history. For example, all members of subfamily II are able to form homodimers on direct repeat sequence, whereas all members of subfamily III can form homodimers on palindromic elements. Monomeric receptors able to bind unique RGGTCA sequences are found in many places on the tree (subfamilies I, IV, and V, for example). One of the most striking cases is the RXRs that are able to form heterodimers only with members of subfamilies I and IV. The only exception to the rule could be the case of COUP-TF and SVP, which have been described as being able to interact with RXRs (Kliewer et al., 1992; Cooney et al., 1993). Nevertheless, this finding is strongly debated and it is not clear whether this interaction represents a real heterodimer formation or could be viewed more as a classical protein–protein interaction (Butler and Parker, 1995). The only physiologically relevant data in favor of this interaction is the ECR-USP/SVP interaction found in Aedes aegypti (Miura et al., 2002; Zhu et al., 2003). It should be noted that the dimer interface of this complex has not been mapped and thus we cannot be sure that it

represents a real heterodimer, such as RXR-TR. Despite this, it is clear that the DNA binding and dimerization abilities are reasonably well correlated with the evolutionary history, suggesting that the first nuclear receptor was able to bind the RGGTCA core sequence, most probably as a homodimer recognizing direct repeat sequences, and that subsequently there was divergence on this theme following the various gene duplications that gave rise to the present diversity of nuclear receptors. A comparison of the phylogenetic tree with the ligand binding ability of the nuclear receptors reveals a very different picture. The following observations clearly suggest that the evolutionary relationships of nuclear receptors and the nature of their ligands are not correlated: i. the presence of nuclear receptors for different ligands within subfamilies (e.g., TRs and RARs, which bind totally different compounds, are related inside subfamily I); ii. closely related ligands recognized by different receptors (e.g., retinoids, which are ligands for RARs, RXRs, and RORb); and iii. the widespread distribution of orphan receptors in the phylogenetic tree. This situation could be explained by an independent gain of ligand binding capacity during nuclear receptor evolution. This model implies that the first member of the nuclear receptor superfamily was an orphan receptor. In accordance with this notion, nuclear receptor sequences described from early metazoans are mostly orthologs of orphan receptors (Escriva Garcia et al., 1997, 2003; Grasso et al., 2001) with the notable exception of the RXR, whose endogenous ligand is still a matter of controversy (Kostrouch et al., 1998; Wiens et al., 2003). Although this model is not discussed in detail here, it is interesting to briefly mention some of its implications. The C-terminal moiety of the nuclear receptor, which is called the ligand binding domain, is well conserved when compared across the whole superfamily, including orphan receptors for which structural data suggest an absence of ligand (e.g., NURR1; Wang et al., 2003). We know that the LBD carries a transcriptional activation domain that can be turned on by a conformational change that allows coactivator binding. It is clear that this conformational change can be induced not only by ligand binding but also by other mechanisms. For example, it is well known that phosphorylation events in the LBD can induce this conformational change (review: Weigel and Zhang, 1998). Alternatively, protein–protein interactions (such as the one occurring between SF1 and Ptx1 on two SF1

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target genes), pituitary luteinizing hormone beta and Mullerian inhibiting substance may also induce coactivator binding (Tremblay et al., 1999). Indeed, the Evans laboratory has suggested that the LBD exists in an equilibrium between an OFF and an ON conformation and that ligand binding directs this equilibrium in favor of the ON (holo) conformation (Schulman et al., 1996). In conclusion, it is believed that ligand binding is only one of the possible switches and that this equilibrium can also be modified by phosphorylation or protein–protein interactions, among other mechanisms. Thus, it is suggested that the sequence conservation of the LBD is not the result of the conservation of ligand binding activity but is rather the hallmark of the existence of this conformational change in all nuclear receptors. The LBD should in fact be viewed as a domain responsible for an allosteric modification of the receptor allowing the exchange between corepressor and coactivator complexes. Seen in this perspective, it is not surprising that the LBD is conserved in orphan receptors. Recent structural data have shed new light on the ligand binding ability of nuclear receptors. First, there is now substantial evidence that ‘‘real’’ orphan receptors (i.e., nuclear receptors that are not recognized by cognate endogenous ligands) may exist. This is the case for NURR1 and its Drosophila homolog HR38, which harbors a LBD folded in an active conformation but with a putative ligand binding pocket filled by large hydrophobic side chains, such that it is difficult to imagine a ligand occupying the same space (Baker et al., 2003; Wang et al., 2003). Other recent reports suggest that in some

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cases a structural ‘‘ligand’’ (e.g., fatty acid for Drosophila USP or for HNF4) may be present inside the LBD without playing any functional role (Billas et al., 2001; Clayton et al., 2001; Wisely et al., 2002). In other cases (ERRg and LRH-1), empty pockets are found in the nuclear receptor, which nevertheless has an active conformation that can be recognized by coactivators (Greschik et al., 2002; Sablin et al., 2003). Thus, the LBD of orphan nuclear receptors can contain pockets filled with side chain residues, pockets with structural ligands or empty pockets, demonstrating an unanticipated large number of possible states between receptors for well-known hormones and orphan receptors. This may render it difficult to define with precision endogenous ligands for a given receptor. Such a debate is currently raging about the RXR and its pharmacological ligand, 9-cis retinoic acid. All these structural results may have evolutionary implications. It has been proposed that the first nuclear receptor had an exogenous molecule such as a fatty acid playing a structural role buried inside its LBD (Sladek, 2002). This compound cannot be called a ligand as it cannot regulate the activity of the receptor and cannot even exit the LBD. A receptor enclosing such a molecule could thus be considered as an orphan receptor such as the Drosophila USP. Second, some of these orphan receptors may have weakened the interaction between these structural ligands and the LBD and ‘‘invented’’ ligand regulated transcriptional activity (Figure 4). Although tempting, this model cannot be generalized and is not widely accepted at present, as recent structural data have shown that LBDs with an empty pocket or with no

Figure 4 A possible scheme for the evolution of ligand binding abilities of nuclear receptors. The original nuclear receptor may have been able to bind a lipophilic compound that was a ‘‘structural ligand,’’ i.e., an integral component of the protein structure. In contrast, modern ligands are interchangeable elements that induce allosteric functional changes of the LBD. This model may explain why the LBD has been conserved during evolution. NR, nuclear receptor; CoR, corepressor; CoA, coactivator. Helix H12 is shown as a yellow box, the ligand as a red box, and the structural ligand as an irregular yellow form.

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pocket are possible and apparently stable. The existence of ‘‘structural ligands’’ is thus perhaps less frequent than anticipated. 3.6.2.2. Metazoan Phylogeny and Nuclear Receptors: the Ecdysozoan Problem

p0085

3.6.2.2.1. Distribution of nuclear receptors in metazoans Recent molecular data have considerably refreshed our ideas and concepts on metazoan phylogeny (Aguinaldo et al., 1997; de Rosa et al., 1999). Figure 5 depicts the present consensus on metazoan phylogeny. Although lively debates are ongoing on many aspects of this tree, several points should nevertheless be briefly discussed in relation to nuclear receptor evolution. The focus will be on the main metazoan phyla since for the vast majority of the 35 phyla no sequence information was available except some classical phylogenetic markers such as mitochondrial genes or ribosomal rRNA genes. 1. The identity of the unicellular eukaryote that is most closely related to metazoans has long been a mystery, but it is now clear that choanoflagellates, which share several cytological and molecular synapomorphies with metazoans, are the sister group of metazoans. Despite some ancient claims based on cross hybridization data or poor sequence identities, it is clear that nuclear receptors are not found outside the Metazoa. There is no known nuclear receptor homolog in the various plant or fungi genomes available and since more than a dozen such genomes are now available, the hypothesis of a secondary loss can be

Figure 5 Simplified phylogeny of metazoans. Clades where NRs have been isolated are labeled with , clades where complete genome sequences exist without any NR gene are labeled with , and clades where the presence of NRs is not yet known are labeled with ? (Modified from Aguinaldo, A.M., Turbeville, J.M., Linford, L.S., Rivera, M.C., Garey, J.R., et al., 1997. Evidence for a clade of nematodes, arthropods and other moulting animals. Nature 387, 489–493; and Adoutte, A., Balavoine, G., Lartillot, N., de Rosa, R., 1999. Animal evolution: the end of the intermediate taxa? Trends Genet. 15, 104–108.)

excluded. But this information is not sufficient to conclude that nuclear receptors are specific to metazoans. Indeed, nothing is known about the presence of nuclear receptors in choanoflagellates. Expressed sequence tag (EST) projects in choanoflagellates have not revealed nuclear receptors, but these genes are usually not frequent in these data because of a relatively low level of expression (King et al., 2003). Indeed, the isolation of a nuclear receptor from the sister group of the Metazoa could illuminate our views on nuclear receptor origins, which remains obscure. Several reports have suggested low scores of sequence identity for the LBD and DBD regions of nuclear receptors with a peroxisomal membrane protein, Pex11p, and the LIM/ GATA zinc finger domain, respectively. This has prompted some groups to suggest that the first nuclear receptor arose from the fusion of two genes encoding these various proteins (Clarke and Berg, 1998; Barnett et al., 2000). In the absence of structural data that can confirm the significance of these low similarity scores, it is hard to draw firm conclusions on this matter. The question of the origin of the first nuclear receptor is thus still open. 2. The metazoans are monophyletic and sponges (Porifera) are clearly the first group that diverged from the metazoan trees. It is now believed that the sponges are not a monophyletic group and the precise phylogeny of the early metazoan tree is far from being resolved but the basal position of sponges when compared to cnidarians is widely accepted. Cnidarians (sea anemone, corals, Hydra) and other related groups such as the Ctenophora diverged later on. The presence of nuclear receptors in sponges has long been debated. In a polymerase chain reaction (PCR) screen, we found nuclear receptor signatures in all animal phyla analyzed from diploblastic animals (e.g., cnidarians; Escriva Garcia et al., 1997) to vertebrates, but not in sponges. Recently, however, Werner Muller’s group reported the presence of an RXR ortholog in the sponge Suberites domuncula (Wiens et al., 2003). Even if the precise identity of this receptor is unclear, it is undoubtedly a member of subfamily II that currently contains, to the exclusion of the RXR, only orphan receptors and which appears to be the most ancient one. The ancestry of subfamily II has been clearly established in our PCR screen that yields subfamily II homologs (COUP-TF, RXR) together with a subfamily V homolog (FTZ-F1) in Hydra and Anemonia, two cnidarians (Escriva Garcia et al.,

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1997). This has been confirmed by the isolation of several nuclear receptors (RXR (NR2B), TR2/4 (NR2C), TLL (NR2E), COUP-TF (NR2F)) in various cnidarians such as Montipora verrucosa, Tripedalia cystophora, and Acropora millepora (Kostrouch et al., 1998; Grasso et al., 2001). Of note, is the presence of subfamily V members in cnidarians, which has not been confirmed by independent evidence and should thus be regarded as tentative. Thus, only subfamily II is clearly present in all metazoans, suggesting that the first nuclear receptor and thus the origin of the superfamily probably belonged to this subfamily. 3. The bilaterians, metazoans that possess three embryonic layers and bilateral symmetry, are split in two main groups, the protostomians and deuterostomians, which differ on many morphological and molecular characters. Interestingly, all bilaterians are now included in one of these two groups. Protostomians are split into two stable groups: the ecdysozoans (animals that molt) including arthropods and nematodes; and the lophotrochozoans including mollusks, annelids, and flatworms (Aguinaldo et al., 1997). Several complete genomes of members of the Ecdysozoa are available (Caenorhabditis elegans, Caenorhabditis briggsae, Drosophila melanogaster, Anopheles gambiae), while the lophotrochozoans are a genomic desert with very few data available and no complete genome sequence available as yet. Thus, many surprises are expected to come from the molecular and comparative genomic study of this group. Indeed, the comprehensive phylogenetic analysis of all nuclear receptors found in complete genome sequences, as well as the identification of an ER ortholog in a mollusk, strongly suggest that liganded receptors such as TRs, RARs, or SRs that were believed to be specific to the vertebrate lineage are probably more widespread among metazoan animals than previously thought (Thornton et al., 2003; Bertrand et al., in press). Importantly, analysis of the complete nuclear receptor set in a given genome allows one to establish unambiguously whether this genome has lost some nuclear receptor if these genes are present in other metazoan species. For example, the fact that Drosophila, C. elegans, and Ciona genomes do not harbor an ER may be viewed as an argument in favor of the fact that ERs are found only in recent chordates, even if the topology of the tree indicates an ancestry of ER in bilaterians. But the recent cloning of a clear ER ortholog in a mollusk (even if this receptor apparently does

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not bind estrogens!) permits reconciliation of the tree topology with the model and leads to the suggestion that indeed an ER ortholog was present in the common ancestor of all bilaterians and was lost independently in nematodes and the Drosophila genomes as well as in Ciona. This example illustrates very clearly the derived nature of the Drosophila and nematode genomes and the importance of lophotrochozoan analysis. Thus, the search for nuclear receptor genes in lophotrochozoans is clearly a priority for future research. The analysis of all nuclear receptors present in bilaterians allows us to propose the basic set of nuclear receptors in metazoan phyla. The principle of this inference is that for each group of nuclear receptors we can locate the speciation event that led to the deuterostome and protostome lineages by the divergence between chordate, insect, and nematode orthologs. Then all genes resulting from duplications, which occurred before this speciation, were probably present in Urbilateria. Using this reasoning, which remains correct even if secondary loss occurred in some lineages, it appears that the genome of the common ancestor to nematodes, insects, and chordates had at least 25 nuclear receptor genes (Bertrand et al., in press). Any variation from this number should be explained by lineage specific events such as gene loss (i.e., arthropods and nematodes) or gene duplications that arose often in certain lineages (i.e., vertebrates or nematodes). 3.6.2.2.2. Nuclear receptors, nematodes, and the ecdysozoan The case of ecdysozoans is of course particularly relevant for the analysis of nuclear receptor evolution in arthropods. Nematodes, arthropods, and other phyla such as onychophorans are grouped in the Ecdysozoa, animals that share the developmental trait of molting, although this grouping is still debated (Aguinaldo et al., 1997; Blair et al., 2002). Genomic data reveal that nematodes contain a very large number of nuclear receptor genes (up to 270) whereas insects (Drosophila and Anopheles) harbor 21 nuclear receptor genes. Three questions are relevant for nuclear receptor evolution in ecdysozoans: (1) what was the extent of gene loss events that occurred during ecdysozoan evolution; (2) in this context, by what mechanisms did nematodes reach such an impressive number of nuclear receptors and did gene duplication play any role in nuclear receptor evolution in arthropods; and (3) are nuclear receptors linked to the grouping of ecdysozoans and their molting behavior? As discussed above, the common ancestor of all bilaterians probably had 25 nuclear receptor genes in its genome. Since Diptera genomes contain 21

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genes, a simple conclusion could be that arthropods lost four genes. In fact, the phylogenetic analysis of the nuclear receptors of bilaterians reveals a much more complex and interesting picture (Bertrand et al., in press). Both Drosophila and nematode genomes lost several genes such as TR, RAR, PPAR, ER, SR, EAR2, and LXR. Whether these genes have been lost only once in the common ancestor of Drosophila and nematodes (i.e., the ancestor of all Ecdysozoa) or twice independently in these two lineages remains unclear. We will need information on other ecdysozoans to resolve this issue. In addition, it is clear that nematodes (or at least C. elegans) lost other genes such as HR39, ERR, RXR, TLL, DSF, PNR, EcR, and E75. It is possible that some of these genes still exist in nematode genomes but that their assignation is obscured by strong sequence divergence since nematode nuclear receptors genes, like many other genes, evolved rapidly (Robinson-Rechavi et al., unpublished data). It is of course very interesting to see that the analysis of dipteran genomes in the context of other bilaterians reveals the loss of seven nuclear receptor genes among which six are encoded liganded receptors. It is important to note, however, that this conclusion is reached only by the analysis of the nuclear receptor set present in the Drosophila and Anopheles complete genomes. There is a long road from the common ancestor of Ecdysozoa to the common ancestor of Diptera and it is not absolutely clear when these losses occurred and whether they occurred simultaneously. Thus, we can expect that some of these genes will be found in some ecdysozoans and even in some arthropods or primitive insects. Since many researchers working on arthropods have been influenced by the set of nuclear receptors present in Drosophila, it came as no surprise that for example SR, ER, or PPARs were not searched for in more primitive insects or arthropods. These data would allow us to revise the nuclear receptor set present in Ecdysozoa. Another basic question suggested by this analysis is why arthropods lost so many liganded receptors and what are the remnants of these signaling pathways still present in the genome of these organisms. This will be the subject of future analysis. Returning to the premise that the urbilateria genome probably had 25 nuclear receptors, of which seven were lost in Ecdysozoa leading to 18 nuclear receptor genes, we then find that dipteran genomes contain 21 nuclear receptor genes. Interestingly, the three supplementary receptors are the three highly unusual members of the KNI group (knirps, knirps-related, and eagle) that clearly form a group of relatively recent duplicates. In nematodes, the

situation is much more striking since the nematode genome lost more genes (7 þ 8 ¼ 15) but contains more than 270 receptors. The phylogenetic analysis of nematode nuclear receptors led to show that the C. elegans genome contains 13 ancient nuclear receptors and approximately 250 supplementary receptors that are the result of an explosive diversification of a unique gene, HNF4. In fact, it can be concluded that C. elegans as well as C. briggsae and probably others contain approximately 250 HNF4 genes! These genes are highly divergent and not functionally or structurally linked and the reason for such a massive increase in gene number is still far from being clear (Robinson-Rechavi et al., unpublished data). Nevertheless, it should be emphasized that examples of lineage specific expansion in gene number are found for other types of genes and in other genomes suggesting that this is an event more widespread than expected. In the perspective of nuclear receptor evolution, it is fundamental to recall, given the important role played by EcR and several other nuclear receptors, that the ecdysozoan group is composed of animals that molt. In this context, it is tempting to speculate that the ecdysone signaling cascade, including the EcR-USP heterodimer would be an obvious ecdysozoan synapomorphy (de Mendonca et al., 1999). It is thus striking, and even irritating, to find neither an ecdysone receptor nor an RXR in the C. elegans genome. Did molting evolve several times independently in Ecdysozoa? The lack of resolution of this issue illustrates once again that we should be very cautious in the conclusions that we draw from classical model organisms such as C. elegans or Drosophila. Indeed, if C. elegans effectively lacks EcR and USP homologs, this unexpected property is not widespread in nematodes. Dirofilaria immitis, a parasitic nematode, does contain an ortholog of EcR and USP (Shea et al., 2003). Thus, before concluding that a given receptor is absent in a large zoological group such as nematodes one should always also look at data from several species not too closely related. Unfortunately, this striking situation also suggests that C. elegans will be a poor model for comparing events governing molting in nematodes with the same events in arthropods. The other question related to the ecdysozoan issue is whether the molting and EcR-USP signaling cascades are widespread outside the Ecdysozoa. If the answer is yes, then this no longer can be viewed as an ecdysozoan synapomorphy. The lack of resolution of this question underlines the profound ignorance of molecular mechanisms regulating the development in protostomians (besides C. elegans and D. melanogaster) in general and in lophotrochozoans in

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particular. Ecdysteroids have been described in Schistosoma but the significance of this observation is far from clear (de Mendonc¸ a et al., 2000a). To date, no EcR ortholog has been found in Schistosoma, which is an interesting observation given that a very large number of EST sequences, including many nuclear receptors, are available in this species (VerjovskiAlmeida et al., 2003). Thus, with our present, very limited knowledge, the proposal that EcR-USP signaling represents an ecdysozoan synapomorphy, although probably oversimplistic, is still acceptable.

3.6.3. Phylogeny of Nuclear Receptors in Arthropods 3.6.3.1. Towards an Inventory

Despite their functional importance, the evolution of arthropod nuclear receptors has been largely overlooked (Henrich and Brown, 1995). Three reasons may account for this disappointing situation. First, the sequencing effort has focused mainly on Diptera (Drosophila, malaria vector An. gambiae, yellow fever mosquito Ae. aegypti) and Lepidoptera (silk worm B. mori, tobacco hornworm M. sexta), which are two closely related and highly derived holometabolous orders of insects. Therefore, the sample of available sequences is small and evolutionarily incomplete (Figure 6). Second, these proteins have not been studied in insects as a homogeneous family of transcription factors, such as the products of Hox genes for example. Rather, they were analyzed separately through their involvement in various developmental pathways, like embryonic segmentation (knirps, tailless, Ftz-F1), molting and metamorphosis (EcR, Usp, Ftz-F1, E75. . .), or eye morphogenesis (seven-up). However, their common structure and mode of action requires an integrated view of their evolution. Third, to the exclusion of EcR, ligands still remain to be identified for the nuclear receptors of arthropods. Whether they are all real orphans or not is still an open question and an essential issue is to understand their function and their evolution. Nevertheless, we can draw here a summary of the present knowledge, in order to suggest the orientations of the future work that should help to understand both the evolution of nuclear receptors and their role during diversification of insects and other arthropods. All the arthropod nuclear receptors known to date can be classified into the seven subfamilies previously described (Figure 1) (Laudet, 1997). For many of them it is easy to identify a sequence relationship with a chordate receptor. However, it appears that rapid evolutionary divergence produced nuclear

Figure 6 Phylogeny of identified nuclear receptors in arthropods. The schematic unrooted tree of the six subfamilies is a simplified version of that from the Nuclear Receptors Nomenclature Committee (1999). In addition, the NR0 subfamily is shown at the bottom of the figure. Branches in bold are those leading to a subfamily with the corresponding number written above the branch. Two names are given for each nuclear receptor: on the left is the official nomenclature and on the right is the trivial name used in this chapter. The number of species where one given receptor has been identified (update: July 2003) is also indicated. Four groups of species have been defined: D, Diptera; L, Lepidoptera; I, other insects; and A, other arthropods (Bonneton et al., 2003). When a nuclear receptor is found in all of these four groups, its name is underlined.

receptors that may be specific to insects or other arthropods. Because ECR and USP-RXR show such a divergence and have been intensively studied (see Chapter 3.5), their evolution will be considered separately. 3.6.3.1.1. Subfamily 1: all members of the ecdysone pathway In insects, subfamily NR1 contains five receptors, which are all directly involved in the ecdysone pathway of D. melanogaster (Figure 7). They can be separated into two distinct groups, based on structural as well as functional data (Laudet, 1997; Thornton and Desalle, 2000). The first group includes E75 (NR1D3), E78 (NR1E1), and HR3 (NR1F4). All of their vertebrate homologs

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f0035

Figure 7 Phylogenetic distribution of identified nuclear receptors in arthropods. On the left, a tree indicates the evolutionary relationships between the groups where nuclear receptors are known (Kristensen, 1981; Whiting et al., 1997; Hwang et al., 2001; Giribet et al., 2001). Diptera is written in bold with two asterisks () to highlight the fact that two complete genome sequences are available for this order (D. melanogaster and An. gambiae), allowing for an accurate account of nuclear receptors (Bertrand et al., in press). Concerning Hymenoptera, we have found HR3, HNF4, SVP, and ERR in Apis ESTs. The 16 groups of arthropod nuclear receptors are indicated by rectangular boxes colored according to their subfamilies, which are shown at the bottom of the figure (subfamilies 0, I, II, III, IV, V, VI). The ongoing complete genome (G) and EST sequencing projects are listed on the right (GOLD); –, no known project.

are orphan receptors that are unable to form heterodimers with RXR. The second group contains the famous ecdysone receptor, EcR (NR1H1), and the less well-known HR96 (NR1J1). Both proteins may be specific to Ecdysozoa. Note that several nuclear receptors of subfamily 1 found in chordates have no homologs in arthropods and nematodes: thyroid hormone receptors (TRs, NR1A), retinoic acid receptors (RARs, NR1B), and peroxisome proliferator-activated receptors (PPAR, NR1C) that bind fatty acids (Laudet and Gronemeyer, 2002). In fact, a recent analysis of data provided by genomic projects suggests that this group of ‘‘classical’’ hormone receptors may be very ancient (present in the ancestor of bilateria) and was lost in ecdysozoans (Bertrand et al., in press).

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3.6.3.1.1.1. E75 The E75 gene (Eip75B in the FlyBase, NR1D3) has been identified in D. melanogaster (Feigl et al., 1989; Segraves and Hogness, 1990), An. gambiae (Bertrand et al., in press), Ae.

aegypti (Pierceall et al., 1999), and several species of Lepidoptera (Segraves and Woldin, 1993; Jindra et al., 1994a; Palli et al., 1995; Palli et al., 1997b; Zhou et al., 1998; Swevers et al., 2002). It was also isolated from the shrimp Metapeneus ensis (Chan, 1998). These proteins are homologous to the vertebrate receptors REVERB-a and REVERB-b, which have been shown to be constitutive transcriptional repressors involved in the pacemaker controlling the circadian clock in vertebrates (Preitner et al., 2002) (see Chapter 4.11). In insects, E75 acts as a repressor of HR3, probably through direct interaction, both in Drosophila (White et al., 1997) and in Bombyx (Swevers et al., 2002). The possible implication of E75 in circadian rhythmicity is still not known (see Chapter 4.11). In Drosophila, inactivation of all E75 functions causes first instar larval lethality, but isoform specific null mutations reveal different functions for each of the three isoforms (Bialecki et al., 2002). The complex role of this gene is far from being fully understood but expression

Evolution of Nuclear Hormone Receptors in Insects

and hormonal induction data suggest that its involvement in early ecdysone responses during molting and metamorphosis may be shared among arthropods (Jindra et al., 1994a; Palli et al., 1997b; Chan, 1998; Zhou et al., 1998). It also plays a role during oogenesis and vitellogenesis in Drosophila (Bryant et al., 1999), Aedes (Pierceall et al., 1999; Raikhel et al., 2002) (see Chapter 3.9), and Bombyx (Swevers et al., 2002). 3.6.3.1.1.2. E78 The E78 gene (Eip78C in the FlyBase, NR1E1) is only known in Diptera (Drosophila and Anopheles) and although it is related to E75 and REVERB, it is extremely divergent from both of them. It could be a paralog of E75 that experienced a rapid evolutionary rate. Interestingly, the same situation is observed in nematodes, with the presence of an ortholog of E75 (nhr-85 in C. elegans) together with an ortholog of E78 (SEX-1 in C. elegans) (Kostrouch et al., 1995; Carmi et al., 1998; Unnasch et al., 1999; Sluder and Maina, 2001; Crossgrove et al., 2002). It is therefore possible that, in addition to one NR1D gene related to REVERB, all ecdysozoans also possess one specific fast-evolving NR1E gene that has been lost in chordates (Bertrand et al., in press). However, mutant phenotypes indicate that this NR1E gene probably plays different roles in insects and in nematodes. Indeed, in Drosophila, E78 homozygous mutants are viable and fertile, with subtle defects in regulation of some puffs and in formation of dorsal chorionic appendages (Russel et al., 1996; Bryant et al., 1999). By contrast, the E78 ortholog (SEX-1) of Caenorhabditis is required for sex determination and viability of hermaphrodites (Carmi et al., 1998). 3.6.3.1.1.3. HR3 Orthologs of HR3 (Hr46 in FlyBase, NR1F4) have been identified in Diptera (Koelle et al., 1992; Kapitskaya et al., 2000; Bertrand et al., in press) and Lepidoptera (Palli et al., 1992; Jindra et al., 1994b; Palli et al., 1995, 1996; Matsuoka and Fujiwara, 2000; Debernard et al., 2001), and a short sequence (partial DBD) is also available from the coleopteran Tenebrio molitor (Mouillet et al., 1999). This receptor is well conserved outside insects, with one gene in Caenorhabditis (Kostrouch et al., 1995) and three homologs in vertebrates: ROR a, b, g, which are transcriptional activators (Jetten et al., 2001). It has been shown recently that cholesterol and all-trans retinoic acid are ligands for RORa (Kallen et al., 2002) and RORb (Stehlin-Gaon et al., 2003), respectively. RORs and REVERB bind to the same response element and RORs are thought to be competitors for REVERB. As such, they are believed to play an important role in circadian rhythm (Andre

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et al., 1998). HR3 is an ecdysone inducible early–late gene that plays a key role during Drosophila metamorphosis by repressing early genes and directly inducing the prepupal regulator FTZ-F1 (Lam et al., 1997; White et al., 1997). This function in molting and metamorphosis is probably conserved throughout insects, as suggested by studies of expression and hormonal induction in Manduca (Palli et al., 1992; Lan et al., 1997, 1999; Langelan et al., 2000), Choristoneura fumiferana (Palli et al., 1996, 1997a), and Tenebrio (Mouillet et al., 1999). Interestingly, inhibition of HR3 expression using RNAi showed that this gene is also required for Caenorhabditis molting, suggesting a conserved role in Ecdysozoa (Kostrouchova et al., 1998, 2001). As for E75, HR3 is a functionally complex gene involved in many aspects of the ecdysone pathway. For example, a role in oogenesis and vitellogenesis has been described for Aedes (Kapitskaya et al., 2000; Li et al., 2000; Raikhel et al., 2002) (see Chapter 3.9), Bombyx (Eystathioy et al., 2001), and Caenorhabditis (Kostrouchova et al., 1998). HR3 is also expressed in embryos of Drosophila (Carney et al., 1997) and Caenorhabditis (Kostrouchova et al., 1998). Furthermore, Drosophila HR3 mutants have defects in their nervous system and die during embryogenesis (Carney et al., 1997). Given the wide variety of tissues and stages of expression for HR3, it is clear that other functions will be identified by detailed analysis of mutants. This is nicely illustrated by two studies showing a specific role for HR3 during the formation of wings in Drosophila and in Bombyx. Clonal analysis reveals requirements for Drosophila HR3 in the development of adult wings, bristles, and cuticle, but no apparent function in eye or leg development (Lam et al., 1999). In a Bombyx wing deficient mutant called flu¨gellos, HR3 expression is reduced only in wing discs, and not in the testis and fat body, while ECR, USP, E75, and HR38 are not affected (Matsuoka and Fujiwara, 2000). The wings are one of many organs where HR3 is expressed, and the role of this gene in their development in two different insects could only be revealed with appropriate genetic manipulations. 3.6.3.1.1.4. HR96 The HR96 (NR1J1) gene has been identified in Drosophila (Fisk and Thummel, 1995) and in Anopheles (Bertrand et al., in press). In Caenorhabditis, three homologs are known: DAF-12, NHR-8, and NHR-48 (Sluder and Maina, 2001). All these ecdysozoan receptors are related to the vertebrate receptors NR1I: VDR, which is the receptor for vitamin D, PXR and CAR, which both bind a wide variety of xenobiotics (Laudet, 1997). Given the fact that the vertebrate homologs of

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HR96 bind steroid-derived compounds, it is likely that HR96 also binds such a ligand. Since ECR and its vertebrate homologs LXR and FXR can bind steroids, it is even tempting to speculate that all the related nuclear receptors NR1H/J have the same property. It is also possible that HR96 requires USP-RXR to bind DNA in the same way as the other NR1H/I/J receptors. Little is known about the functions of HR96, which is inducible by 20E and is expressed during the onset of metamorphosis (Fisk and Thummel, 1995). The Caenorhabditis homologous gene daf-12, which functions downstream of the insulin and TGF-b signaling pathways, regulates diapause, developmental age, and adult longevity (Antebi et al., 1998, 2000; Gerisch et al., 2001; Jia et al., 2002). Interestingly, a growing number of studies suggest that common regulatory mechanisms control developmental timing in Caenorhabditis and Drosophila, with a central role for the ecdysone pathway in insects (Thummel, 2001) (see Chapters 3.3 and 3.5). s0080

3.6.3.1.2. Subfamily 2: a large collection of old and new genes Only subfamilies NR2 and NR5 have been found in all metazoans, suggesting that they are probably at the origin of the superfamily (Escriva et al., in press). Most of the NR2 proteins are orphans (except HNF4 and RXR) and do not form heterodimers (except USP-RXR). In insects, subfamily NR2 contains eight genes, including the most conserved nuclear receptors: HNF4, seven-up (SVP-COUP), and tailless (TLL). The tailless group (NR2E) contains four different genes that exhibit a rather restricted pattern of expression: TLL, PNR, DSF, and FAX1. Probably they all share a primary function in the developing nervous system (see Chapter 2.4). 3.6.3.1.2.1. HNF4 HNF4 (NR2A) is one of the best-conserved nuclear receptors between arthropods and vertebrates. This gene was identified in Drosophila (Zhong et al., 1993), Aedes (Kapitskaya et al., 1998), Anopheles (Bertrand et al., in press), and Bombyx (Swevers and Iatrou, 1998). Two homologous HNF4 genes exist in mammals. In Caenorhabditis, a surprising explosive burst of duplications of HNF4 produced most of the (>270) supplementary nuclear receptors found in this species (Robinson-Rechavi et al., unpublished data). The embryonic expression pattern of this receptor is remarkably well conserved between insects and vertebrates. Indeed, HNF4 is transcribed zygotically in several analogous organs: the midgut, Malpighian tubules, and fat body of insects (Zhong et al., 1993;

Kapitskaya et al., 1998; Swevers and Iatrou, 1998) and the intestine, kidney, and liver of vertebrates (Laudet and Gronemeyer, 2002). Furthermore, analyses of mutants show that the development of these organs requires HNF4 activity in Drosophila (Zhong et al., 1993) as well as in the mouse (Chen et al., 1994). If conflicting results were obtained concerning the role of HNF4 in the fat body of Drosophila embryos (Zhong et al., 1993; Hoshizaki et al., 1994), this gene is clearly transcribed in the fat body of Aedes adults (Kapitskaya et al., 1998) as well as in the Bombyx larvae and pupae (Swevers and Iatrou, 1998). In the mosquito, three isoforms have been identified, each having a specific temporal pattern during the vitellogenesis that takes place in the female fat body (Kapitskaya et al., 1998). Therefore, HNF4 probably performs similar functions during the formation of the gut and its derivatives in a large range of metazoans. Another conserved pattern is the maternal expression of HNF4, but its function has not been studied. It has been shown recently that the mammalian HNF4 receptor binds fatty acids constitutively (Dhe-Paganon et al., 2002; Wisely et al., 2002). Given the strong similarity (85%) observed for the HNF4 LBD between insects and vertebrates, it is tempting to speculate that this type of ligand may also be used in insects. 3.6.3.1.2.2. HR78 The HR78 gene (NR2D1) has been identified in Drosophila (Fisk and Thummel, 1995; Zelhof et al., 1995b), Anopheles (Bertrand et al., in press), Bombyx (Hirai et al., 2002), and Tenebrio (Mouillet et al., 1999). These genes are distantly related to the vertebrate’s orphan receptors TR2 and TR4, and to the very divergent nematode genes nhr-41 (Caenorhabditis; Sluder and Maina, 2001) and nhr-2 (filarial nematode Brugia malayi; Moore and Devaney, 1999). As with most of the early genes of the ecdysone cascade (including ECR, E75, and E78), the developmental expression of HR78 is globally stable between the species of the D. melanogaster subgroup (Rifkin et al., 2003). HR78 is required for ecdysteroid signaling during the onset of metamorphosis of Drosophila (Fisk and Thummel, 1995; Zelhof et al., 1995b; Fisk and Thummel, 1998). This receptor is inducible by 20E and binds to over 100 sites on polytene chromosomes, many of which correspond to ecdysteroid regulated puff loci. By contrast with the important sequence divergence of the proteins, the temporal profile of expression of HR78 is very well conserved between Drosophila and Tenebrio, with an early activation at the end of larval stages, followed by a rather constant level during prepupal stages (Fisk

Evolution of Nuclear Hormone Receptors in Insects

and Thummel, 1995; Zelhof et al., 1995b; Mouillet et al., 1999). Drosophila HR78 null mutants die during the second and third larval instar with tracheal defects, showing that, despite a uniform expression in the embryo, this receptor is not essential for early development (Fisk and Thummel, 1998; Astle et al., 2003). A surprising result was obtained by studying the Bombyx homolog. Indeed, yeast two hybrid and pull down assays showed that Bombyx HR78 could interact with USP-RXR (Hirai et al., 2002). If we consider that HR78 and ECR bind to the same DNA sites (Fisk and Thummel, 1995; Zelhof et al., 1995b), and have the same USP-RXR partner, it is therefore possible that competition occurs between these two proteins during the onset of metamorphosis. 3.6.3.1.2.3. SVP The seven-up gene (SVP, NR2F3) is a member of the COUP-TFs group, the most conserved nuclear receptors. It has been identified in various insects: Drosophila (Henrich et al., 1990; Mlodzik et al., 1990), Aedes (Miura et al., 2002), Bombyx (Togawa et al., 2001), and partial clones were also recovered from Tenebrio (Mouillet et al., 1999) and the grasshopper Schistocerca gregaria (Broadus and Doe, 1995). One homolog was studied in Caenorhabditis (Zhou and Walthall, 1998), and three COUP-TF genes are present in vertebrates (Pereira et al., 2000). These orphan receptors have a very broad pattern of expression and are essential for development in all the organisms studied. In the embryos of Drosophila and Schistocerca, SVP was found in the developing eyes, the central and peripheral nervous systems, the Malpighian tubules, and the fat body (Mlodzik et al., 1990; Hoshikazi et al., 1994; Broadus and Doe, 1995; Kerber et al., 1998; Sudarsan et al., 2002). Analysis of mutants demonstrated the functional role of these expressions in Drosophila. In addition, SVP participates in the diversification of cardioblasts in the heart tube of Drosophila embryos (Lo and Frasch, 2001; Ponzielli et al., 2002). The SVP protein is regulated by different pathways, such as Ras in eyes, EGF in Malpighian tubules, and hedgehog in heart, and thus plays multiple roles from cell fate determination to regulation of cell cycle and differentiation. Interestingly, SVP/COUP-TF inhibits USP-RXR signaling pathways, both in vertebrates and in insects. This has been tested in vivo with Drosophila in which the overexpression of SVP that leads to lethality at the onset of metamorphosis can be rescued by a concomitant overexpression of USP-RXR (Zelhof et al., 1995a). It is of note that SVP is expressed at the beginning of metamorpho-

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sis, when the ecdysteroid titer is low, both in Drosophila and in Tenebrio (Mouillet et al., 1999). A similar inhibition of 20E response occurs during vitellogenesis in the mosquito Aedes (see Chapter 3.9). Furthermore, in this species, it was shown in vitro that the repression of the ecdysone pathway is probably due to a direct interaction between SVP and USP-RXR. The replacement of ECR by SVP in USP-RXR heterodimers would be essential for the termination of vitellogenesis, when the ecdysteroid titer declines (Miura et al., 2002; Zhu et al., 2003). However, it must be noted that no physical interaction between SVP and USP-RXR was detected in Drosophila (Zelhof et al., 1995a). 3.6.3.1.2.4. TLL The tailless gene (TLL, NR2E2) is one of the most conserved nuclear receptors. It was first identified in Drosophila as a terminal gap gene determining embryo segmentation (Ju¨ rgens et al., 1984). Homologs have been studied in Drosophila virilis (Liaw and Lengyel, 1993), the house fly Musca domestica (Sommer and Tautz, 1991), and the coleopteran Tribolium castaneum (Schro¨ der et al., 2000). Nematodes and vertebrates also have one tailless gene. The early terminal expression is necessary for the establishment of the nonmetameric domains at the anterior and posterior poles of the Drosophila embryo (Ju¨ rgens et al., 1984; Pignoni et al., 1990). This pattern is very well conserved in Diptera (Sommer and Tautz, 1991; Liaw and Lengyel, 1993). By contrast, the early posterior expression of tailless in Tribolium reveals a temporal divergence. This difference suggests that tailless may not function as a gap gene in Tribolium, but may be involved in an earlier specification of terminal fate (Schro¨ der et al., 2000). Thus, it appears that an important shift occurred in tailless function during the transition from short-germ to long-germ embryogenesis. This situation contrasts with the conservation of tailless late expression in the developing forebrain of insects and vertebrates. Using mutants demonstrated an essential role for tailless in eye formation of Drosophila (Daniel et al., 1999; Hartmann et al., 2001) and the mouse (Monaghan et al., 1997; Yu et al., 2000). In conclusion, the primary conserved function for tailless would be in the development of the forebrain, while its role in segmentation was probably acquired during the evolution of long-germ holometabolous insects. Other gap genes such as orthodenticle, empty spiracles, or hunchback are known to be part of a conserved neural network that was recruited for insect segmentation (Reichert, 2002).

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3.6.3.1.2.5. PNR One NR2E3 gene, homologous to the vertebrate photoreceptor-specific nuclear receptor (PNR; Kobayashi et al., 1999), was identified in the genome of Drosophila (Flybase: CG16801) and Anopheles (Bertrand et al., in press). Nothing is known about this gene in insects. It is of note that PNR expression is restricted to the retina and plays a critical role in the development of photoreceptors (Kobayashi et al., 1999; Haider et al., 2000). Therefore, both tailless and PNR play an important role during vertebrate eye development. We have seen that the role of tailless in the formation of the visual system is conserved between insects and vertebrates. Whether the same conservation exists for PNR remains to be established. 3.6.3.1.2.6. DSF The NR2E4 nuclear receptor (dissatisfaction, DSF) may be specific to insects. It has been identified in D. melanogaster (Finley et al., 1998), D. virilis (Pitman et al., 2002), Anopheles (Bertrand et al., in press), and Manduca (Pitman et al., 2002), but no homologs are found in nematodes and chordates. Recent analysis of genomic data suggest that DSF is one of the four nuclear receptors (along with HR39, E78, and FAX1) that were lost in the chordate lineage (Bertrand et al., in press). DSF is necessary for appropriate sexual behavior and sex-specific neural development. It acts downstream of the genes Sex lethal and transformer in the sex determination cascade. Mutant females resist male courtship and fail to lay eggs, while mutant males are bisexual and mate poorly (Finley et al., 1997; O’Kane and Asztalos, 1999). The dsf gene is expressed in a very limited set of neurons in the brain of larvae, pupae, and adults, with no sex specificity (Finley et al., 1998). The DSF protein acts as a repressor, through its LBD and an unusually very long hinge region (Pitman et al., 2002). It will be very interesting to test whether DSF can also act as a ligand-dependent activator, since no sex hormones are known in insects (De Loof and Huybrechts, 1998).

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3.6.3.1.2.7. FAX1 The NR2E5 gene, called FAX1, was first identified in Caenorhabditis as a regulator of axon pathfinding and neurotransmitter expression (Wightman et al., 1997). FAX-1 is expressed in embryonic neurons after neurogenesis but just prior to axon extension (Much et al., 2000). Homologs were identified in the genome of Drosophila (Flybase: CG10296) and Anopheles (Bertrand et al., in press). The fax-1 genes probably experienced a rapid evolution in Ecdysozoa, since they are very divergent from all the other genes of the NR2E group of nuclear receptors.

3.6.3.1.3. Subfamily 3: ERR a classical steroid receptor in insects? The subfamily NR3 comprises the receptors for sex and adrenal steroid hormones: estrogens (ER), androgens (AR), progesterone (PR), glucocorticoids (GR), and mineralocorticoids (MR), as well as ERR which is an orphan receptor related to ER. Until recently, these proteins were believed to be restricted to vertebrates. However, not only is one ERR gene present in the urochordate Ciona intestinalis (Yagi et al., 2003) and in Diptera, but an ER ortholog was isolated from a mollusk (Thornton et al., 2003). Therefore, it appears that, to the exclusion of ERR, steroid receptors from the subfamily NR3 were specifically lost in ecdysozoans (Bertrand et al., in press). Unfortunately, the precise relationships between ER, ERR, and the other steroid receptors are difficult to determine with the current data. One ERR gene (NR3B4) is present in the genome ¨ stberg et al., of Drosophila (Adams et al., 2000; O 2003) and Anopheles (Bertrand et al., in press). It was also found independently in a screen for Drosophila embryonic neural precursor genes (Brody et al., 2002). There is no ortholog of ERR in the Caenorhabditis genome (Maglisch et al., 2001). In vertebrates, the three ERR genes have broad expression patterns and influence a wide range of physiological processes, including those governed by ERs (Giguere, 2002). A striking contrast between vertebrate and insect endocrinology is the lack of sex determining hormones in insects (De Loof and Huybrechts, 1998). Obtaining functional data for Drosophila and/or Anopheles ERR could help to tackle this intriguing problem. 3.6.3.1.4. Subfamily 4: HR38, an alternative partner for USP-RXR The subfamily NR4 is a small group of orphan nuclear receptors: the vertebrate’s NGFIB a bg, divergent nematode receptors (NHR-6 in C. elegans) and the insect HR38 (Sluder and Maina, 2001; Laudet and Gronemeyer, 2002). The HR38 gene (NR4A4) has been cloned in Drosophila (Fisk and Thummel, 1995; Sutherland et al., 1995; Komonyi et al., 1998), Aedes (Zhu et al., 2000), and Bombyx (Sutherland et al., 1995). Remarkably, like their vertebrate homologs NGFIB, insect HR38 can bind DNA either as a monomer or through an interaction with USPRXR (Sutherland et al., 1995; Crispi et al., 1998; Zhu et al., 2000). This interaction plays a very interesting role in the mosquito Aedes. In this insect, vitellogenesis requires a blood meal that triggers a 20E cascade to activate yolk synthesis (see Chapter 3.9). Before a blood meal, this regulation is inhibited despite the presence of the hormone.

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Evolution of Nuclear Hormone Receptors in Insects

A key factor of this inhibition is the competitive binding of HR38 to USP-RXR, which prevents the formation of the ECR/USP-RXR heterodimer, the functional EcR (Zhu et al., 2000; Raikhel et al., 2002). Therefore, although HR38 is not directly regulated by 20E, it can participate in the 20E pathway as an alternative partner to USP-RXR. In Drosophila, HR38 is expressed in ovaries and during all stages of development. Different mutant alleles have different lethal phases, from larval stages to adults, showing a role in metamorphosis and adult epidermis formation (Kozlova et al., 1998). Recent results have shown that the Drosophila heterodimer HR38/USP-RXR responds to a distinct class of ecdysteroids independently of ECR and without direct binding of the ligand to either HR38 or USP-RXR. In fact, crystal structure analysis reveals that the Drosophila HR38 LBD lacks both a conventional ligand binding pocket and a bona fide AF2 transactivation domain (Baker et al., 2003). Interestingly, the vertebrate NGFIB receptors are ligand independent transcriptional activators that are considered to be real orphans, a view which was recently reinforced by structural analysis (Wang et al., 2003). These data provide strong evidence that NR4 receptors operate through an atypical mechanism of activation. 3.6.3.1.5. Subfamily 5: competition in the ecdysone pathway The subfamily NR5 contains orphan nuclear receptors that, together with NR2, are believed to be at the origin of the superfamily (Escriva et al., in press). In mammals, the subfamily NR5 contains three homologs, including the steroidogenic factor 1 (SF1), an essential regulator of endocrine function (Parker et al., 2002). One ortholog (nhr-25) is present in the genome of Caenorhabditis (Gissendanner and Sluder, 2000). In insects, the two NR5 genes, FTZ-F1 and HR39, are important in the ecdysone (20E) pathway.

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3.6.3.1.5.1. FTZ-F1 The FTZ-F1 gene (NR5A3) is one of the best-known nuclear receptors of insects. This gene encodes two protein isoforms, a and b, each having a specific role. The aFTZ-F1 isoform acts during segmentation of the early embryo, a function that is likely to be an innovation of holometabolous insects. The bFTZ-F1 isoform is a major regulator of metamorphosis, a role probably shared by all ecdysozoans. Interestingly, FTZ-F1 has been cloned not only in Diptera (Lavorgna et al., 1991, 1993; Li et al., 2000) and Lepidoptera (Sun et al., 1994; Weller et al., 2001), but also in the honeybee, Apis mellifera (Hepperle and Hartfelder, 2001), Tenebrio (Mouillet et al., 1999), and in the

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shrimp Metapeneus ensis (Chan and Chan, 1999). Only ECR and USP-RXR have been identified in such a wide sample of arthropod species (Figure 6). The Drosophila aFTZ-F1 is a direct regulator of the pair-rule gene fushi-tarazu (ftz), which governs the formation of embryonic metameres (Ueda and Hirose, 1990; Lavorgna et al., 1991; Ohno et al., 1994). Later, it was also found that aFTZ-F1 and FTZ are mutually dependent cofactors for regulation of target genes such as engrailed, wingless, or ftz itself (Florence et al., 1997; Guichet et al., 1997; Yu et al., 1997). This interaction requires the DBD of both proteins as well as a contact between a LRALL domain of the FTZ protein with the AF-2 activation domain of aFTZ-F1 (Schwartz et al., 2001; Suzuki et al., 2001; Yussa et al., 2001). The LRALL domain matches the consensus box LxxLL that is found in many cofactors of nuclear receptors (Laudet and Gronemeyer, 2002). Such a partnership between a homeodomain protein and a nuclear receptor is still a unique example in insects (Yussa et al., 2001). It raises the exciting possibility that nuclear receptors may interact with Hox genes. Indeed, ftz is a member of the arthropod Hox gene family. However, it is a very derived Hox gene, which has lost its homeotic function in insects and acquired new roles in neurogenesis and segmentation (Hughes and Kaufman, 2002). Thus, it was interesting to determine whether the LxxLL box was conserved in the FTZ proteins. Elegant experiments, both in vitro and in vivo, revealed a progressive evolution of FTZ in insects. The pairrule function is moderately conserved in T. castaneum and absent in the Schistocerca. Furthermore, this functional evolution correlates with the presence of a LxxLL domain in the FTZ protein of the holometabolous insects Drosophila and Tribolium, and its absence in the hemimetabolous grasshopper Schistocerca (Alonso et al., 2001; Lohr et al., 2001). In conclusion, it appears that the ftz gene gained one of the domains that facilitate the interaction between aFTZ-F1 and FTZ somewhere in the emergence of holometabolous insects. The isoform bFTZ-F1 plays a central role during molting and metamorphosis of Drosophila. It is repressed by 20E and its expression, which occurs during mid-prepupa when the titer of 20E is low, activates the late prepupal genes (Lavorgna et al., 1993; Woodard et al., 1994). Its activation requires HR3 (Kageyama et al., 1997; White et al., 1997), an interaction that is also used during vitellogenesis of Aedes (Li et al., 2000) (see Chapter 3.9). Analysis of mutants shows that bFTZ-F1 provides competence for stage-specific responses to 20E throughout the organism (Broadus et al., 1999; Yamada

304 Evolution of Nuclear Hormone Receptors in Insects

et al., 2000). These general characteristics seem to be well conserved in Lepidoptera, as shown by expression and induction experiments made with Bombyx (Sun et al., 1994) and Manduca (Hiruma and Ridifford, 2001; Weller et al., 2001). In fact, the patterns of expression observed in Tenebrio (Mouillet et al., 1999) and the shrimp M. ensis (Chan and Chan, 1999), together with the mutant analysis of the Caenorhabditis homologous gene nhr-25 (Asahina et al., 2000; Gissendanner and Sluder, 2000) suggest that the role of bFTZ-F1 may be very similar in all Ecdysozoa. A promising result is that bFTZ-F1 could be one of the factors controlling honeybee caste development (see Chapter 3.13). In this social insect, JH and ecdysteroids induce a dramatic reduction in ovariole number during the fourth larval instar of workers but not in queens. In a search for genes controlling this phenotypic plasticity, the bFTZ-F1 of A. mellifera was identified by differential display of RT-PCR as a 20E downregulated gene in the larval ovary (Hepperle and Hartfelder, 2001). This pioneering work highlights one of the most interesting putative functions for nuclear receptors: molecular regulation of polyphenism (Nijhout, 2003) (see Chapter 3.13).

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3.6.3.1.5.2. HR39 The HR39 gene (NR5B1) exists in Drosophila (Ohno and Petkovich, 1992; Ayer et al., 1993), Anopheles (Bertrand et al., in press), and Bombyx (Niimi et al., 1997) but not in vertebrates. Note that this gene was initially described with the misleading name of FTZ-F1b but should not be confused with the b isoform of FTZF1 (NR5A3). This second group of subfamily NR5 was previously seen as the result of a duplication that arose during insect evolution (Laudet and Gronemeyer, 2002). However, the identification of an HR39 homolog in Schistosoma mansoni, a platyhelminth parasite of humans, suggests that the FTZF1/HR39 duplication occurred very early during protostomian evolution (de Mendonc¸ a et al., 2002). The Caenorhabditis genome has one FTZ-F1 gene (nhr25) but no HR39 homolog (Sluder and Maina, 2001). The HR39 gene of Drosophila is induced by 20E and expressed at every stage of development, with a maximum at the end of third larval and prepupal stages (Horner et al., 1995). Therefore, contrary to bFTZ-F1, it behaves like an early gene during the onset of metamorphosis. Remarkably, the pattern of expression of Schistosoma HR39 resembles that of Drosophila, with the highest amounts in the larval form (de Mendonc¸ a et al., 2002). Both HR39 and FTZ-F1 bind as monomers to the same response elements (Ayer and Benyajati, 1992, 1993; Ohno and Petkovich, 1992; Crispi

et al., 1998). Overexpression of HR39 or FTZ-F1 have opposite effects on promoter activity, suggesting competition between these two nuclear receptors (Ayer et al., 1993; Ohno et al., 1994). 3.6.3.1.6. Subfamily 6: HR4, a new early–late gene? The HR4 gene (NR6A2) is homologous to germ cell nuclear factor (GCNF), an orphan nuclear receptor of vertebrates. It has been identified in the genome of Drosophila (Adams et al., 2000) and Anopheles (Bertrand et al., in press), in the lepidopterans Manduca (Weller et al., 2001), Bombyx (Charles et al., 1999), and Trichoplusia ni (Chen et al., 2002) as well as in Tenebrio (Mouillet et al., 1999). One homolog is also found in nematodes (Sluder and Maina, 2001), showing that there might be one single NR6A gene in all metazoans. For once, Drosophila is not the leading model organism in the study of an insect nuclear receptor, since most of our current information on HR4 come from Tenebrio and lepidopteran species. Several lines of evidence suggest that HR4 may be a new early–late gene involved in the onset of metamorphosis. First, HR4 is directly inducible by 20E in Manduca (Hiruma and Riddiford, 2001) and in a Trichoplusia cell line (Chen et al., 2002). Then, its activation during molting and metamorphosis starts after the early gene HR3 and before the mid-prepupal gene FTZ-F1. Manduca, Bombyx, and Tenebrio share this pattern of expression (Charles et al., 1999; Mouillet et al., 1999; Hiruma and Riddiford, 2001; Weller et al., 2001). Finally, preliminary results obtained with Drosophila confirm that HR4 plays an essential role during molting and metamorphosis (King-Jones and Thummel, FlyBase). However, as for many other nuclear receptors, the most conserved functions are probably in embryogenesis and gametogenesis. Indeed, in adult vertebrates, GCNF is predominantly expressed in the germ cells of gonads, and loss of GCNF function causes embryonic lethality (Chung and Cooney, 2001). Furthermore, the Drosophila HR4 gene was also identified in an enhancer-trap screen for novel X-linked essential genes. The P-insertion (PL78) located in this gene causes embryonic lethality with observable cuticular defects and reveals that Drosophila HR4 is expressed in embryonic gut and ovary (Bourbon et al., 2002). 3.6.3.1.7. Subfamily 0: atypical transcription factors KNI, KNRL, and EG The subfamily 0 was artificially created to encompass all the proteins that contain only one of the two conserved domains (DBD: NR0A or LBD: NR0B) of the nuclear receptors (nuclear receptors nomenclature committee, 1999). In insects, three transcription factors have

Evolution of Nuclear Hormone Receptors in Insects

been classified in the subfamily NR0A because they lack a LBD but contain a DBD similar to the one found in the superfamily (Laudet, 1997). This group contains three genes that have been cloned only in Diptera: knirps (KNI, NR0A1), knirps-related (KNRL, NR0A2), and eagle (EG, NR0A3). No homolog can be found in nematodes and vertebrates, suggesting that this group may be specific to insects or arthropods. On phylogeny based with the DBD, these genes are distantly related to HR96 and ECR, typical nuclear receptors of subfamily NR1 (Laudet, 1997). The gap gene knirps has been identified in D. melanogaster (Nauber et al., 1988), D. virilis (Gerwin et al., 1994), D. americana, D. novamexicana (Wittkopp et al., 2003), and Musca (Sommer and Tautz, 1991) but, surprisingly, it is absent from the genome of Anopheles (Bertrand et al., in press). KNRL and EG are present both in Drosophila and Anopheles (Oro et al., 1988; Rothe et al., 1989; Higashijima et al., 1996; Bertrand et al., in press). The chromosomal location of these genes, at the proximal part of the left arm of the third chromosome, is conserved between Drosophila and Anopheles. These data suggest that knirps may be the result of a duplication of an ancestral knrl gene, which occurred during the evolution of brachyceran Diptera. This hypothesis is reinforced by the strong conservation of expression and function between Drosophila knirps and knrl. Indeed, both genes are functionally redundant for their role in determining the anterior head structures at the blastoderm stage (Gonzalez-Gaitan et al., 1994). Furthermore, they control together the development of several other organs, such as trachea (Chen et al., 1998), wing veins (Lunde et al., 1998, 2003), foregut, and hindgut (Fuss et al., 2001). Thus, it appears that knirps and knrl, which are very close together on the third chromosome, have conserved the same regulatory regions after duplication. An interesting difference, however, is the size of their intron sequences: one kb in knirps and 19 kb in knrl. The consequence of this difference in intron size is that knrl can complement knirps mutations only as an artificial intronless transgene. This effect is explained by the required coordination of mitotic cycle length and gene size during the rapid developmental period of gap gene expression (Rothe et al., 1992). The eagle gene (initially named egon, for embryonic gonads) is required for the specification of serotonergic neurons and other neuroblasts in the embryonic and larval central nervous system of Drosophila (Higashijima et al., 1996; Dittrich et al., 1997; Lundell and Hirsch, 1998). This gene is present in

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the genome of Anopheles (Bertrand et al., in press), but nothing is known about eagle in other species of insects. 3.6.3.2. Conservations and Innovations

What general conclusions can be drawn from our present knowledge of the evolution of nuclear receptors in insects? First, it is likely that all insects have at least 17 different nuclear receptors, but DSF, KNI, KNRL, and EG are not included in this number, since their origin by domain recombination is poorly understood. They might well be insectspecific duplications, but this hypothesis has yet to be tested. There are 21 nuclear receptor genes in the genomes of both the brachyceran dipteran Drosophila (Adams et al., 2000) and the nematoceran dipteran Anopheles (Bertrand et al., in press). In other insects, given the absence of complete genomic sequences, the full number of these genes is not yet known. However, thanks to a wide interest in the genetics of metamorphosis, several nuclear receptors have been cloned by classical means outside Diptera. In Lepidoptera, 12 nuclear receptors have been identified (11 in B. mori), eight in Coleoptera (seven in Tenebrio molitor) seven in Hymenoptera and only three in Orthoptera (Figure 7). Different ongoing genome and EST sequencing projects should help to clarify this fundamental point (GOLD). Indeed, among the model organisms chosen for these projects are representatives of Hymenoptera (the honeybee A. mellifera), Coleoptera (the corn rootworm D. virgifera), crustaceans (the water flea Daphnia, the crab C. pugilator, and the shrimp F. chinensis), and Chelicerates (the tick Amblyomma americanum). Despite the current lack of heterometabolous and ametabolous insects in these projects, the sequence data will be essential to draw a comprehensive phylogenetic analysis of arthropod nuclear receptors. With respect to functional evolution, three major trends can be seen (Figure 8). First, a strong conservation of structure and regulation characterize HNF4, SVP, and TLL. These proteins are the most conserved nuclear receptors in metazoans and although they may have acquired new functions in insects, their fundamental roles in patterning the embryo are likely to be very similar in all species. There is little doubt that HNF4, SVP, and TLL genes will be found in all insects and other arthropods. Second, by contrast, KNI, KNRL, and EG show an extreme divergence, with the lack of a LBD. In that respect, they cannot be considered as bona fide nuclear receptors. Third, it appears that most of the other nuclear receptors have a broad pattern of expression and participate, directly or indirectly, in

306 Evolution of Nuclear Hormone Receptors in Insects

Figure 8 Developmental characteristics of insect nuclear receptors. The tree is the same as in Figure 6. The column on the left (mutant) indicates the stage of lethality of the D. melanogaster mutants: E, embryo; L, larva; L1, first instar larva; L3, third instar larva; P, pupa; A, adult. In addition, the viable phenotypes (V) are also indicated. The central column (pattern) summarizes the spatial and temporal patterns of expression in insects as broad (B) or restricted (R). The column on the right (ecdysone) indicates whether one given nuclear receptor is part of the ecdysone pathway (+) or simply interacts with it ().

the 20E pathway (Figure 8) (Sullivan and Thummel, 2003). It is a general characteristic of nuclear receptors to be expressed at various stages in many different organs, therefore participating in several physiological processes. The organism uses them as powerful gene regulators whose activity can be coordinated in time and space through the availability of the appropriate ligand. The current view may be biased by the fact that the single hormone mostly related to the insect’s nuclear receptor is 20E. However, this signal has undoubtedly acquired a central role in the developmental timing controlled by nuclear receptors.

3.6.4. Evolution of the Ecdysone (20E) Receptor 3.6.4.1. The Arthropod Ecdysone Receptor is a Heterodimer between ECR and USP-RXR

Within the superfamily of nuclear receptors, ECR (NR1H1) belongs to the same group as the vertebrate liver X receptors (LXRa and LXRb__NR1H3, and NR1H2_), which are receptors for oxysterols (Laudet and Gronemeyer, 2002). USP-RXR (NR2B4) is the ortholog of vertebrate retinoid X receptors (RXRa, b, .g NR2B1, 2, 3) (Laudet and Gronemeyer, 2002). The name USP (ultraspiracle)

Evolution of Nuclear Hormone Receptors in Insects

comes from the phenotype of Drosophila mutants (Perrimon et al., 1985), whereas RXR refers to the mammalian ligand (9-cis retinoic acid) (Mangelsdorf et al., 1990). Both genes have not only been isolated in a large number of dipteran and lepidopteran species, but also in several other insect species and arthropods (Figure 6). The functional Drosophila ecdysone (20E) receptor is a heterodimer of the products of the ecdysone receptor (EcR) and ultraspiracle (usp) genes (Koelle et al., 1991; Oro et al., 1992; Yao et al., 1993). The requirement of heterodimerization between ECR and USP-RXR has been found in all the other species studied, including several dipterans (Wang et al., 1998; Vo¨ gtli et al., 1999), lepidopterans (Swevers et al., 1996; Perera et al., 1998; Lan et al., 1999; Minakuchi et al., 2003), the crab C. pugilator (Durica et al., 2002), and even the tick Am. americanum (Guo et al., 1997, 1998). Therefore, it appears that the heterodimer ECR/USP-RXR is the functional ecdysone (20E) receptor in all arthropods. It must be noted that these tests of heterodimerization were made using artificial or real Drosophila EcREs, and that the natural binding sites are unknown in other species. However, in Aedes (Wang et al., 1998), as well as in Drosophila (Vo¨ gtli et al., 1998), the heterodimer ECR/USPRXR exhibits a broad DNA binding specificity, which is likely to be conserved among arthropods. 3.6.4.2. Divergence of the Ecdysone Receptor in Diptera and Lepidoptera

Understanding the evolution of ecdysteroid regulation in insects requires comparative analysis of both partners of the heterodimer. It has been shown that ECR and USP-RXR experienced a strong acceleration of evolutionary rate in Diptera and Lepidoptera compared with other insects (Bonneton et al., 2003). In the LBD of USP-RXR there is only 49% identity between Diptera–Lepidoptera and other insects, as opposed to 68% between these other insects and other arthropods, and 70% between

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the other insects and chordates (Table 1). Thus, the USP-RXR LBD of many insects is less similar to Diptera and Lepidoptera than it is to the chordate RXRs. A similar situation holds for the DBD and LBD domains of ECR, although the divergence is less pronounced. Thus, both ECR and USP-RXR LBD sequences of Diptera and Lepidoptera have evolved at significantly different rates than other species (Table 1). This acceleration defines a clear separation within holometabolous insects. Diptera and Lepidoptera belong to the clade Panorpida (Kristensen, 1981), with Hymenoptera as a sister group. Panorpida also includes: Trichoptera (caddisflies), the sister group of Lepidoptera, as well as Mecoptera (scorpionflies), and Siphonaptera (fleas) which are more closely related to Diptera (Figure 6). A unique event of acceleration could be responsible for the accelerated evolutionary rates at the base of and within these groups. 3.6.4.3. Evolution of USP-RXR

Several protein domains for which sequence divergence is specific to Diptera and Lepidoptera have been identified (Figure 9). The most impressive differences affects the LBD of USP-RXR. In addition, ECR sequences show variability in other domains, namely the DNA binding and the C-terminal F domains (Bonneton et al., 2003). In the Drosophila and Heliothis USP-RXR structures, the loop between helices H1 and H3 is located inside the hydrophobic furrow of the LBD, thereby preventing the repositioning of helix H12 and interactions with coactivators, and locking these USP-RXRs in an unusual antagonist conformation (Billas et al., 2001; Clayton et al., 2001). In the light of these results, our observation of Diptera- and Lepidoptera-specific sequence diversity in both the loop H1–H3 and the helix H12 suggests a form of concerted evolution between these two interacting regions of the USP-RXR LBD. This evolution may have changed the ligand-dependent transactivation activity of

Table 1 Identity and evolutionary rates for USP-RXR and ECR LBD USP-RXR LBD Groups of species

Diptera-Lepidoptera/ Diptera-Lepidoptera/ Other insects/

Identity (%)

Other insects Other arthropods Other arthropods

49 44 68

ECR LBD Evolutionary rate 

0.31 0.36 0.06 NS

Identity (%)

Evolutionary rate

64 58 68

0.12 0.13 0.006 NS

Values indicated for comparisons of evolutionary rates are substitution rate difference. The probability associated to the test is indicated as follows: not significant (NS) > 5%;   5%;   0.5%. (Adapted from Bonneton, F., Zelus, D., Iwema, T., RobinsonRechavi, M., Laudet, V., 2003. Rapid divergence of the ecdysone receptor in Diptera and Lepidoptera suggests coevolution between ECR and USP-RXR. Mol. Biol. Evol. 20, 541–553.)

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Figure 9 Protein divergence and functional domains of ECR and USP-RXR in insects. On this diagram (not to scale), the DBDs are shown in black, the LBDs in white, and the Diptera-Lepidoptera specific insertions are in grey. Dashed lines indicate regions with a strong Diptera/Lepidoptera divergence. The N-terminal A/B domain (not shown) and the linker region (D domain) are highly divergent in size and sequences and, thus, are not compared in this figure. The average identity percentages of pairwise comparisons for DBD and LBD of USP-RXR and ECR are from Bonneton et al. (2003). The name of each divergent domain is indicated, with its known or putative function written below.

the protein. It may also have had an effect on the ligand binding activity, since the loop H1–H3 contains residues that interact with the phospholipid cocrystallized with Drosophila and Heliothis LBD. However, given the very strong conservation of Helix H10, it is likely that the dimerization activity of USP-RXR LBD remained largely unchanged during evolution. This is important, since USPRXR is the heterodimeric partner of numerous nuclear receptors, both in vertebrates (TR, RAR, PPAR, LXR, FXR, CAR, NGFIB, etc.) and in insects (HR38, SVP, HR78). All these hypotheses can now be tested both in vitro and, more importantly, in vivo using transgenesis. Small changes at the molecular level can lead to important functional differences that may be revealed only by studying their consequences in whole organisms. For example, replacing the domains D and E of the Drosophila usp gene with the equivalent domains of another dipteran, Chironomus, allows the rescue of usp mutants from first instar larvae lethality, but do not restore a normal metamorphosis (Henrich et al., 2000). Therefore, despite the 72% of similarity between the exchanged domains, this chimeric USP-RXR fails to function normally in Drosophila. Similar studies involving smaller chimeric regions and site directed mutations should help in the

future to identify which domains contributed to the functional evolution of USP-RXR in insects. 3.6.4.4. Evolution of ECR

It is intriguing that the LBD of ECR underwent a significant increase of substitution rate in Diptera and Lepidoptera, while its structure remained apparently largely unchanged. In all insects, and presumably in all arthropods, ECR LBD binds 20E (Riddiford et al., 2000). This fundamental interaction may represent the primary selective constraint acting on this domain. However, the ligand affinity of ECR can show subtle differences in insects. This is best illustrated within Diptera. A homology model of the Chironomus ECR LBD showed that the putative ligand binding pocket is strongly conserved in all arthropods (Wurtz et al., 2000). Nevertheless, comparative tests of toxicity and receptor affinity of nonsteroid insecticides revealed that the Chironomus ecdysone (20E) receptor behaves more like a lepidopteran than like Drosophila (Smagghe et al., 2002). Also of great interest is the discovery that the Aedes receptor is significantly much more sensitive to ecdysone than the Drosophila receptor, whereas both proteins exhibit the same sensitivity to 20E, the natural biologically active hormone. Surprisingly, the genetic origin of this difference in

Evolution of Nuclear Hormone Receptors in Insects

ligand specificity was mapped into the helix H10 of the ECR LBD, a region known to be essential for dimerization (Wang et al., 2000). These results show that the conservation of the structure of ECR LBD can hide a significant plasticity, which can only be revealed through appropriate functional tests. Nuclear receptor LBDs are also involved in heterodimerization activity. The rapid evolution of ECR can be explained by adaptation to the extremely divergent USP-RXR, and eventually acquisition of new partners, such as HR38 (Baker et al., 2003). It may be that the stability of the heterodimer required compensatory changes in ECR and USPRXR, suggestive of coevolution. The differences seen in ECR DBD also suggest functional changes in dimerization (Bonneton et al., 2003). Indeed, four of the six substitutions that are conserved among Diptera and Lepidoptera are located in the second zinc finger, at positions known to be involved in protein dimerization but not in DNA contact or nuclear localization signal (Black et al., 2001; Khorasanizadeh and Rastinejad, 2001). Another evolutionary acquisition of Diptera and Lepidoptera is the presence of a C-terminal F domain of variable length, which does not show any sequence conservation between species (Figure 8; Bonneton et al., 2003). Most nuclear receptors do not contain this domain, including mammalian homologs LXRs. This difference is interesting, since it is known that when present (ERa, HNF-4) the F domain of nuclear receptors can modulate different functions of the LBD (Montano et al., 1995; Nichols et al., 1998; Peters and Khan, 1999). The F domain of Drosophila ECR may also contribute modestly to the activation function of the LBD (Thornmeyer et al., 1999; Hu et al., 2003). 3.6.4.5. Ecdysone Receptor and the Evolution of Insects

Ecdysteroids control reproduction and development of arthropods. As a result, the pattern of expression of the ecdysone (20E) receptor correlates very well in time and space with the localization and the level of these hormones. Indeed, both ECR and USP-RXR are expressed in a wide range of tissues during oogenesis, embryogenesis, molting, and metamorphosis (see Chapter 3.5). Therefore, any evolutionary change in the mode of action of the ecdysone receptor can have important consequences for development and/or reproduction. This major issue has only started to be addressed, and most of the questions are still unanswered. Changing the regulation of a key developmental gene is probably one of the most effective ways to create new functions rapidly during the course

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of evolution. In that respect, very interesting results have shown that alterations in the timing of ovarian ECR and USP-RXR expression characterize the evolution of larval reproduction in two species of gall midges (Hodin and Riddiford, 2000). In these dipterans, pedogenesis involves the precocious growth and differentiation of the ovary and subsequent parthenogenetic reproduction in an otherwise larval form. Genetic analysis has shown that ECR and USP-RXR regulate the timing of ovarian morphogenesis during Drosophila metamorphosis (Hodin and Riddiford, 1998). These data suggest that heterochronic shifts in the ovarian regulation of both genes may play an important role in the evolution of dipteran pedogenesis. In a larger perspective, a growing number of evidence suggest that the genetic cascade triggered by ecdysteroids may play a central role in the control of developmental timing in Ecdysozoa (Thummel, 2001). Therefore, we can expect that future studies will identify changes in the expression of the ecdysone receptor associated with the evolution of different temporally regulated processes of insects such as, for example, molting, metamorphosis, diapause, and adult longevity. Regarding the evolution of postembryonic development, it is observed that the divergence of EcR does not correlate with the different types of insect metamorphosis (Bonneton et al., 2003). Moreover, several nuclear receptors such as E75, HR3, HR78, bFTZ-F1, and HR4 play very similar roles in the control of molting and metamorphosis amongst insects (see Chapter 3.5). These results suggest that the ecdysteroid-induced genetic cascade (at least the upstream part of this cascade) is a well-conserved developmental module in insects; this conclusion may even be extended to all the molting metazoans. Actually, theories that try to explain the evolution of insect metamorphosis usually consider that JH is the key player, rather than 20E (Sehnal et al., 1996; Truman and Riddiford, 1999, 2002). A heterochronic shift in embryonic JH secretion, with an earlier appearance of this hormone, could well have been the major event in the transition from hemimetabolous to holometabolous insects. In the light of these interpretations, the possibility that JH is a natural ligand of USP-RXR (Jones and Sharp, 1997; Jones et al., 2001; Sasorith et al., unpublished data) is one of the most important point to clarify in order to understand the emergence of metamorphosis in insects.

Acknowledgments We thank Barbara Demeinex and Hector Escriva for their very helpful comments on this text, and Gilles

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Parmentier for the search of nuclear receptors in honeybee’s EST. Work from our laboratory is funded by the CNRS, the MENRT, the Re´ gion Rhoˆ ne-Alpes, and the ARC.

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3.7 The Juvenile Hormones W G Goodman, University of Wisconsin–Madison, Madison, WI, USA N A Granger, University of North Carolina, Chapel Hill, NC, USA ß 2005, Elsevier BV. All Rights Reserved.

3.7.1. Introduction 3.7.2. Chemistry of the Juvenile Hormones 3.7.2.1. Discovery of the Major Juvenile Hormone Homologs 3.7.2.2. Hydroxylated Juvenile Hormones 3.7.3. Other Naturally Occurring Juvenile Hormones 3.7.3.1. Metabolites of Juvenile Hormones as Hormones? 3.7.3.2. Other Juvenile Hormones 3.7.4. Biosynthetic Pathways for the Juvenile Hormones 3.7.4.1. Specificity of the Biosynthetic Pathways 3.7.4.2. Possible Feedback Loops in the Biosynthetic Pathways 3.7.4.3. What Constitutes a Juvenile Hormone Titer? 3.7.5. Measurement of the Juvenile Hormones 3.7.5.1. Bioassays 3.7.5.2. Physicochemical Assays 3.7.5.3. Radioimmunoassays 3.7.5.4. Radiochemical Assays 3.7.6. Evolution of the Juvenile Hormones 3.7.6.1. Juvenile Hormone Homolog Distribution among the Insect Orders 3.7.6.2. Evolutionary Connections among the Arthropods 3.7.6.3. Evolutionary Inferences 3.7.7. Regulation of Juvenile Hormone Biosynthesis 3.7.7.1. Neuropeptides 3.7.7.2. Allatostatins 3.7.7.3. Other Factors Regulating Juvenile Hormone Biosynthesis 3.7.7.4. Second Messengers 3.7.7.5. Putting It All Together 3.7.8. Hemolymph Transport Proteins for the Juvenile Hormones 3.7.8.1. High Affinity, High Molecular Weight Hemolymph Juvenile Hormone Binding Proteins 3.7.8.2. High Affinity, Low Molecular Weight Hemolymph Juvenile Hormone Binding Proteins 3.7.8.3. Functions 3.7.9. Catabolism of the Juvenile Hormones 3.7.9.1. Juvenile Hormone Esterases 3.7.9.2. Juvenile Hormone Epoxide Hydrolases 3.7.9.3. Secondary Metabolism of Juvenile Hormone: Juvenile Hormone Diol Kinase 3.7.9.4. Juvenile Hormone Catabolism and New Directions 3.7.10. Juvenile Hormones in Embryological Development 3.7.10.1. Juvenile Hormone Homologs and Precursors Present during Embryogenesis 3.7.10.2. Role of Methyl Farnesoate during Embryogenesis 3.7.10.3. Juvenile Hormone Titers during Embryogenesis: Correlation with Developmental Events 3.7.10.4. Roles of Juvenile Hormone during Embryogenesis 3.7.10.5. Juvenile Hormone Binding Proteins of the Embryo 3.7.10.6. Juvenile Hormones and the Evolution of Insect Metamorphosis 3.7.11. Juvenile Hormones in Premetamorphic Development 3.7.11.1. Juvenile Hormone Titers 3.7.11.2. Potential Problems with Titer Determinations 3.7.11.3. Premetamorphic Roles of the Juvenile Hormones

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3.7.12. Molecular Mode of Action of the Juvenile Hormones 3.7.12.1. Juvenile Hormone Interaction with Ultraspiracle 3.7.12.2. Juvenile Hormone Interaction with the Methoprene-Tolerant Gene 3.7.12.3. Juvenile Hormone and Metamorphosis: The Role of the Broad-Complex 3.7.12.4. Juvenile Hormone Regulation of Cytosolic Malate Dehydrogenase

3.7.1. Introduction The juvenile hormones (JHs) represent a family of acyclic sesquiterpenoids that are, with a single known exception, unique to the class Insecta. They are the principal products of the corpora allata (CA), retrocerebral paired or unpaired endocrine glands of ectodermal origin (Wigglesworth, 1970). One or more JHs have been identified in approximately 100 insect species spanning at least 10 insect orders, from the most ancestral to the most highly derived. From an evolutionary standpoint, the JHs may have originally been involved in orchestration of reproductive processes such as control of gonadal development and vitellogenin synthesis. The role of JH in the more highly derived insect orders has expanded considerably to include regulation of metamorphosis, caste determination, behavior, diapause, and various polyphenisms (Nijhout, 1994). This chapter will present the current status of our knowledge about the JHs, their evolution, roles in embryological and larval development, biosynthesis, transport, catabolism, and molecular mode of action. The role of JH in reproduction is covered elsewhere (see Chapter 3.9).

3.7.2. Chemistry of the Juvenile Hormones 3.7.2.1. Discovery of the Major Juvenile Hormone Homologs

Elucidation of the chemical structures of the JHs began with Ro¨ller and colleagues, who identified the principal JH in lipid extracts of Hyalophora cecropia (Ro¨ller et al., 1967). This first JH, methyl (2E,6E 10-cis)-10,11-epoxy-7-ethyl-3,11-dimethyl2, 6-tridecadienoate, was termed JH I, and since that time, eight molecules with similar aliphatic sesquiterpene structures have been identified in insects (Figure 1). The structure was confirmed as the 2E,6E, 10 cis isomer (Dahm et al., 1968), while the absolute configuration of JH I at its chiral centers (C10 and C11) was determined to be 10R,11S (Faulkner and Petersen, 1971; Nakanishi et al., 1971; Meyer et al., 1971). Meyer et al. (1968) subsequently identified a minor component in the H. cecropia extracts that differed from JH I by a methyl, instead of an ethyl

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group at C7 (Figure 1). Termed JH II, methyl (2E, 6E 10-cis)-10,11-epoxy-3,7,11-trimethyl-2,6-tridecadienoate), like JH I, displays an E,E configuration at C2, C3 and C6, C7. The absolute configuration at the C10, C11 positions for the naturally occurring JH II has not been rigorously determined (Baker, 1990); however, in binding studies, (10R,11S)-JH II bound with nearly the same affinity to the Manduca sexta hemolymph juvenile hormone binding protein as (10R,11S)-JH I (Park et al., 1993). Thus, it is likely that the naturally occurring enantiomers of JH I and II display the same 10R,11S configuration. The third JH homolog, JH III, methyl 10,11 epoxyfarnesoate, was identified from medium in which CA of the tobacco hornworm, M. sexta, had been maintained (Judy et al., 1973) (Figure 1). JH III differs from the higher homologs in that the three branches of the carbon skeleton at C3, C7, and C11 are methyl groups; however, it displays the same 2E,6E geometry. The hormone has only one chiral carbon, C10, which, in the naturally occurring hormone, displays the 10R configuration. JH III is the only, or predominant, JH in all insects except Lepidoptera (Schooley et al., 1984). Interestingly, JH III had been chemically synthesized a decade earlier, before it was recognized as a naturally occurring hormone (Bowers et al., 1965). A fourth and fifth JH (Figure 1), the trihomosesquiterpenoids JH 0 and its isomer, 4-methyl JH I (iso-JH 0), were identified in M. sexta eggs (Bergot et al., 1981a), but nothing is currently known of their functions. More recently, JH III with a second epoxide substitution at C6,C7 was isolated and identified from in vitro cultures of larval ring glands of Drosophila melanogaster (Richard et al., 1989) (Figure 1). This homolog, termed JH III bisepoxy (JHB3), is active in a D. melanogaster bioassay and has been found in other higher Diptera (Borovsky et al., 1994a; Moshitzky and Applebaum, 1995; Yin et al., 1995). 3.7.2.2. Hydroxylated Juvenile Hormones

Hydroxylation reactions are frequently associated with the inactivation of bioreactive molecules but in certain cases, hydroxylation can actually increase the biological potency of a compound. This important biochemical reaction appears to be involved in

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Figure 1 Chemical structures of the naturally occurring juvenile hormone (JH) homologs, hydroxylated JHs, JH metabolites, and commonly used JH analogs.

the production of a new group of JH homologs, the hydroxylated JHs (HJHs). These HJHs, 4-OH, 8OH, and 12-OH JH III, are synthesized and released by the CA of the African locust, Locusta migratoria (Darrouzet et al., 1997, 1998; Mauchamp et al.,

1999) (Figure 1). The biological activity of synthetic 12-OH JH III was bioassayed utilizing Tenebrio molitor and was found to be 100-fold more active than JH III (Darrouzet et al., 1997). This result may reflect the cross-species nature of the bioassay,

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since the assay using L. migratoria is not sufficiently sensitive (see Section 3.7.5.1). While the biosynthetic pathway for the synthesis of the HJHs has not been determined, two pathways have been suggested: (1) hydroxylation of the JH III structure itself or (2) incorporation of hydroxylated precursors into the JH backbone via the isoprenoid biosynthetic pathway (Darrouzet et al., 1998). A cytochrome P450, CYP4C7, identified in the cockroach Diploptera punctata, can catalyze the formation of 12-OH JH III and (10E)-12-OH farnesol, indicating that hydroxylation of JH III appears to be a primary reaction in the CA (Sutherland et al., 1998). However, there is a sharp increase in the expression of this enzyme when JH is absent or when there is an intrinsic repression of JH synthesis, suggesting that hydroxylation of JH is a mechanism for inactivating the hormones. The pathways of synthesis and the physiological roles of the HJHs remain a fertile area for investigation.

3.7.3. Other Naturally Occurring Juvenile Hormones Davey (2000a), in a thought-provoking essay, posed an intriguing question: ‘‘How many JHs are there?’’ Using a chemical definition, i.e., that JH is a farnesoid molecule secreted by the CA, one can count but a handful. However, if a JH is defined by its biological activity, the JHs could include HJHs, catabolic products, and nontraditional molecules, increasing the number significantly and creating the difficult problem of sorting out which molecules are actually JHs. 3.7.3.1. Metabolites of Juvenile Hormones as Hormones?

An open mind is needed when considering whether metabolites of JH could serve as substrate for conversion to the biologically active hormone or perhaps could act as hormones themselves. A group of molecules that has long been overlooked and frequently relegated to a mere biochemical curiosity are the JH conjugates (Roe and Venkatesh, 1990). Polar metabolites of JH I that are cleaved by glucosidases and sulfatases were identified in flies nearly three decades ago (Yu and Terriere, 1978), and more recent studies hint at the existence of very polar metabolites of JH. A major metabolite resulting from the injection or application of JH II into wandering larvae and prepupae of the cabbage looper, Trichoplusia ni, was found to be an unidentified, water-soluble polar product of the hormone (Kallapur et al., 1996). Identification by radio

high-performance liquid chromatography (HPLC), as well as mass spectrometry, has confirmed that the molecule is a novel polar metabolite and that it is not a JH diol phosphate (M. Roe, personal communication). In vitro studies on the biosynthetic products of the CA from M. sexta and two other lepidopteran species indicate that the glands are synthesizing JH conjugates not recognized by antibodies highly specific for the JHs and their acids; nevertheless, when these compounds are hydrolyzed by esterases, JH III acid is formed (Granger et al., 1995a). This product also appears to exist in the hemolymph, and the results of this study suggest the moiety is either a glucuronide of JH III or an acylglycerol with JH III as the side chain(s). Schooley’s group identified other polar metabolites in the hemolymph of M. sexta as phosphate conjugates of JH I and JH III diol and found that JH I diol phosphate (Figure 1) is the principal end product of JH I metabolism in M. sexta (Halarnkar and Schooley, 1990; Halarnkar et al., 1993). More recently, Maxwell et al. (2002a, 2002b) identified the enzyme responsible for catalyzing the conversion of the JH diol to the JH diol phosphate, JH diol kinase (JHDK), and proposed that cellular JH epoxide hydrolase (JHEH) and JHDK are primarily responsible for the irreversible inactivation of JH early in the last stadium of M. sexta, when the JH titer rapidly decreases (Baker et al., 1987) (see Section 3.7.9). At the present time, there appear to be only two definitive pathways for the catabolism of JH: (1) hydrolysis of the methyl ester of JH by hemolymph esterases, yielding JH acid, and (2) hydration of the epoxide by JHEH, yielding JH diol (Roe and Venkatesh, 1990) (Figures 1 and 2). Can any of these catabolites act as a JH? There is certainly evidence that JH acid can be converted to JH and that it may also act as a hormone. It has been known for a number of years that JH acids are secreted by the CA of M. sexta beginning early in the last larval stadium (Janzen et al., 1991) and that they are apparently the sole product of the CA by the wandering stage in this stadium (Sparagana et al., 1984; Janzen et al., 1991) (Figure 3). JH acid is also detected in the hemolymph of M. sexta (Baker et al., 1987) and of the silkworm Bombyx mori, where, at certain critical stages, the titer of JH acid surpasses that of JH (Niimi and Sakurai, 1997). JH acid is found as a product of the CA in other lepidopterans including the adult male loreyi leafworm, Mythimna loreyi (Ho et al., 1995a, 1995b), the adult male black cutworm, Agrotis ipsilon (Duportets et al., 1998), and the larval tomato moth, Lacanobia oleracea (Audsley et al., 2000).

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Figure 2 Primary routes of JH metabolism. Route I represents hydrolysis of the methyl ester via an esterase followed by hydrolysis of the epoxide ring via an epoxide hydrolase. Route II represents hydrolysis of the epoxide ring via an epoxide hydrolase followed by hydrolysis of the methyl ester via an esterase. The end product in both routes is JH acid diol.

Studies by Bhaskaran and his colleagues suggest that JH acid may be a hormone in its own right. The ability of M. sexta fat body cells to respond to the JH analog methoprene (Figure 1) with enhanced production of yolk protein and its mRNA is acquired after ecdysteroid-initiated commitment to pupation, but it also requires prior exposure to JH II acid or methoprene acid (Ismail et al., 1998). In subsequent work, Ismail et al. (2000) demonstrated that for two metamorphic events, the production of pupal proteins by Verson’s gland, and the loss of the ability of crochet epidermis to produce larval crochets, exposure to JH acid or methoprene acid plus a low dose of RH5992 (an ecdysteroid analog) is required. Use of RH5992 or methoprene acid alone does not induce these changes. Gilbert et al. (2000) recently outlined unpublished experiments by Bhaskaran and colleagues to determine whether JH acids played a role in the acquisition

of metamorphic competence in abdominal rings of first stadium D. melanogaster. Preliminary results indicate that JH acid plus ecdysteroid induce in first instar D. melanogaster both Broad, a transcription factor that appears in response to ecdysteroids early in the last larval stadium (see Section 3.7.12.3), and the adult promoter of alcohol dehydrogenase. However, attempts to repeat this work have been inconclusive, and the results of the previous works have been questioned on technical grounds. 3.7.3.2. Other Juvenile Hormones

It is a curious fact that the JH of Rhodnius prolixus, the hemipteran used by Wigglesworth in his pioneering experiments on the nature of the hormones controlling growth and reproduction (Wigglesworth, 1936), does not correspond to any other known JH (Wyatt and Davey, 1996). Early

324 The Juvenile Hormones

Figure 3 Biosynthesis of JH and JH acid by Manduca sexta corpora allata (CA) in vitro. Biosynthesis of JH I (open circles) and JH I acid (closed circles) by CA in vitro from fifth instars as measured by radioimmunoassay after separation of JH and JH acid by phase partition. Synthesis is expressed as ng JH I radioimmunoassay equivalents. E, the time of larval ecdysis; W, the onset of wandering behavior; P, the time of pupal ecdysis. Each datum point is the mean of 9–12 incubations (
SEM). (Reproduced from Janzen, W.P., Menold, M., Granger, N.A., 1991. Effects of endogenous esterases and an allatostatin on the products of Manduca sexta corpora allata in vitro. Physiol. Entomol. 16, 283–293.)

attempts to identify the hemipteran JHs did not meet with success (Baker et al., 1988), but a more recent report suggests that JH I is the JH of Hemiptera (Numata et al., 1992). Unfortunately, this latter identification seems doubtful in light of the results of Kotaki (1993, 1996). These studies revealed that while the product of the CA of the stink bug, Plautia stali, has a sesquiterpenoid skeleton similar to JH III, the addition of the JH III precursors farnesoic acid or farnesol to incubations of CA stimulates the production of neither JH III nor JHB3, but one that differs chromatographically from any known JH (Kotaki, 1997). Among the possible groups of known, but nontraditional JHs, are the biosynthetic precursors of JH (Figures 4 and 5). Methyl farnesoate (MF) (Figure 1) may be the crustacean form of the insect JH (see Section 3.7.6; Chapter 3.16), but it is also present during embryogenesis in the cockroach Nauphoeta cinerea (Bu¨ rgin and Lanzrein, 1988) (see Section 3.7.10.2) and is synthesized by the embryonic CA of D. punctata (Cusson et al., 1991a) and the adult CA of the dipteran Phormia regina (Yin et al.,

1995). In P. regina, MF alone has limited JH-like biological activity, but appears to work best in a coordinated role together with JH III and JHB3. Farnesoic acid (FA) (Figure 4), another JH precursor, has also been identified as a product of the D. punctata nymphal CA; moreover, it is proposed to have a hormonal or prohormonal function during the latter half of the fourth stadium, when release of JH III ceases. Alternatively, it may be secreted as a by-product of O-methyl transferase activity in the terminal steps of JH III biosynthesis (Yagi et al., 1991). Cusson et al. (1991a) speculate that continued synthesis of FA would allow for the rapid resumption of JH III synthesis when it is needed. Plants possess compounds that exhibit JH-like activity in insect bioassays, and these molecules are considered to play a defensive role. For the most part, these ‘‘phytojuvenoids’’ are structurally distinct from the insect JHs and, with the exception of farnesol, which occurs widely in flowering plants, do not resemble the JH homologs (Bergamasco and Horn, 1983). Curiously, (10R)-JH III and MF have been conclusively identified in the sedges Cyperus iria and C. aromaticus (Toong et al., 1988). Recently, it has been possible to produce JH III and its precursors and to define its biosynthetic pathway in cell suspension cultures of C. iria (Bede et al., 1999, 2001). Given the large amount of JH III present in the plants, one might suspect that it acts as a naturally occurring insect growth regulator to deter feeding. However, it is possible that the plant uses the hormone as a plant growth inhibitor to retard the growth of other nearby species. There exist a few compounds with no structural similarity to the JHs that display hormonal activity. The ecdysteroids top this short list. Ovarian ecdysteroids have long been known to have JH functions in adult females, stimulating the synthesis of vitellogenin in some species, and terminating previtellogenic reproductive diapause and promoting uptake of yolk proteins by oocytes in others, all roles traditionally associated with JH (Hagedorn, 1983; Girardie and Girardie, 1996; Richard et al., 1998) (see Chapter 3.9). There may also exist peptides that mimic the action of JH. In R. prolixus, JH stimulates protein synthesis in the male accessory gland in vitro, a function also supported by an uncharacterized neuropeptide from the brain (Barker and Davey, 1983; Gold and Davey, 1989). In addition, there are reports of peptides stimulating the synthesis of vitellogenin in L. migratoria, a role usually attributed to JH (Girardie and Girardie, 1996; Girardie et al., 1998). Finally, there is the proposal of Davey (2000b) that thyroid hormones may function

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Figure 4 Biosynthetic pathway for JH III. In Lepidoptera, the last two steps in the biosynthetic pathway convert farnesoic acid to JH acid, which is then methylated to form JH III (1). In Orthoptera and Dictyoptera, the last two steps in the biosynthetic pathway convert farnesoic acid to methyl farnesoate, which is then epoxidized to form JH III (2).

in insects. Phenoxy-phenyl compounds such the JH analog fenoxycarb (Figure 1), thyroxine (T4) and triiodothyronine (T3) mimic the effects of JH III in reducing the volume of follicle cells in L. migratoria (Davey and Gordon, 1996; Kim et al., 1999). In

many species, follicle cells respond to JH by altering their cytoskeletal structure, resulting in the appearance of lateral spaces between the cells (patency) via which vitellogenin gains access to the oocyte surface (Davey et al., 1993). T3 binds to the same receptor

326 The Juvenile Hormones

Figure 5 Biosynthetic pathways for JH 0, I, II, and III.

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on locust follicle cells as JH III (Davey and Gordon, 1996; Kim et al., 1999), and T3 immunoreactivity can be found in locust hemolymph (Davey, 2000a, 2000b). Locust food (wheat shoots and bran) was found to contain both T3 and T4, thus suggesting that insect diets could serve as supplementary sources of compounds which act as JH (Davey, 2000b). The fact that T3 apparently binds to a receptor that also binds JH has interesting implications in the search for potential JH receptor(s) (see Section 3.7.12). The increased sensitivity afforded by modern analytical tools should make easier the difficult task of deciphering the biological roles of these new JH-like molecules. Are they biologically relevant or are they the result of promiscuous biochemical reactions that yield end products with no real biological activity? It remains to be determined whether the results of studies using incubations in vitro or homogenates of CA to generate new JH-like molecules under experimental conditions bear any resemblance to the situation in vivo. There is good evidence from vertebrate endocrinology to believe that it is a real problem. A very early study on corticosteroid biosynthesis in the adrenal gland (Heftmann and Mosettig, 1960) lists more than 40 corticosteroids present in the human adrenal glands, even with the limited technology of that time. Of those, only a handful have ever been detected in the blood. A similar situation has been found with the insect ecdysteroids (Rees, 1985) (see Chapter 3.3).

3.7.4. Biosynthetic Pathways for the Juvenile Hormones The biosynthetic pathway for the JHs, now known for many years, has been described in exquisite detail by Schooley and Baker (1985). Briefly, biosynthesis of the most ubiquitous JH, JH III, proceeds by the normal terpenoid pathway: the formation of five-carbon (5C) isoprenoid units from acetate via mevalonic acid, with the sequential head to tail condensation of three 5C units to form farnesyl pyrophosphate (Figure 4). Farnesyl pyrophosphate then undergoes esteratic cleavage to farnesol, which is then oxidized to FA. The last two steps in the biosynthetic pathway diverge depending upon the insect order. In Lepidoptera, a C10,C11 epoxidase converts the FA to the epoxy acid (JH acid), which is then methylated by an O-methyl transferase to form the methyl ester. In Orthoptera and Dictyoptera, the converse appears to be the case: epoxidation follows methylation (Schooley and Baker, 1985; Figure 4).

327

Synthesis of the higher homologs, JH 0, I, and II, appears to be a unique biochemical feature limited to certain insect species (Schooley et al., 1984). These insects can synthesize homoisoprenoid (6C) units from acetate and propionate via 30 -homomevalonate, and from a mixture of these isoprenoid (5C) and homoisoprenoid (6C) units form the respective homologs (Figure 5). Thus, three homoisoprenoid units (ethylmethylallyl pyrophosphates) form the skeleton of JH 0, two homoisoprenoid units and one isoprenoid unit (dimethylallyl pyrophosphate) form JH I, and one homoisoprenoid unit and two isoprenoid units form JH II. With regard to the pathway for JHB3, Moshitzky and Applebaum (1995) have presented evidence that synthesis in D. melanogaster proceeds from FA, which is epoxidized twice, at C6,C7 and C10, C11 to form 6,7;10,11-bisepoxyfarnesoic acid (FABE). FABE is then methylated to yield JHB3 (Figure 6). It should be noted that JH III is not epoxidized to JHB3. 3.7.4.1. Specificity of the Biosynthetic Pathways

The coexistence of multiple JH homologs in a single species at the same developmental stage, together with the discovery of new forms of JH in insects previously thought to have only one, presents an enigma not only as to their functions, but also as to the substrate specificity of the pathways by which they are synthesized and the regulatory mechanisms that effect those steps. Schooley and Baker (1985) concluded that biogenesis of the JH skeleton occurs via the condensation of isoprenoid and homoisoprenoid units and that overwhelming evidence suggests low substrate specificity at most steps in the biosynthetic pathway, even in enzyme preparations from CA, or incubations of CA, from insects known to produce JH III only. This is particularly evident in the two terminal steps, where lack of substrate specificity has been demonstrated in a number of species. For example, the CA of the locust Schistocerca gregaria, which normally synthesize only JH III in vitro, will produce JH I as their sole product if incubated with dihomofarnesoic acid (Schooley et al., 1978a). Schooley and Baker (1985) concluded that the proportions of the precursors (propionate, mevalonate, and homomevalonate) added to incubation medium in which CA are maintained dictate the proportions of the JH homologs produced. The precursor propionate is essential for the formation of the ethyl side chain of JH 0, I and II (Figure 5), and within the CA, propionate is derived principally from the branched-chain amino acids

328 The Juvenile Hormones

Figure 6 Biosynthetic pathway for JHB3.

isoleucine and valine (Brindle et al., 1988). The CA of D. punctata produce only trace amounts of JH II in vitro when propionate is the sole carbon donor, while continuing to produce copious amounts of JH III, the sole naturally occurring JH in this species (Feyereisen and Farnsworth, 1988). The CA of such insects are unable to convert branched-chain amino acids into propionate due to the lack of an active branched-chain amino acid transaminase (Brindle et al., 1987, 1992). Cusson et al. (1996) tested the hypothesis that shifts in the proportions of the different JHs and JH acids released by lepidopteran CA are associated with changes in the activity of the transaminase. They found that the age-related increase in the biosynthesis of JH I acid by adult male CA of the moth Pseudaletia unipuncta was linked to an age-related increase in transaminase activity in CA homogenates. This increase was not observed with female CA, which synthesized more JH II than JH I during this same time period. The authors propose the existence of a factor regulating propionate production in lepidopteran CA. This factor is not

the M. sexta allatostatin (see Section 3.7.7.2) that inhibits JH biosynthesis by female P. unipuncta CA, since this peptide has no effect on the rate of isoleucine metabolism by either male of female glands. Thus, as predicted, the availability of propionate within the CA and the activity of the transaminase has a significant impact on the proportions of JH I and II and their acids, but has no effect on the production of JH III, which is synthesized from the nonbranched isoprenoid units (Figure 5). The specificity and activity of other enzymes may therefore be involved in creating the blend of JH homologs and their acids synthesized by lepidopteran CA. As noted by Schooley et al. (1976) and Cusson et al. (1996), these enzymes may be the isopentenyl diphosphate isomerase and the farnesyl diphosphate synthase that catalyze the condensation of the isoprenoid and homoisoprenoid units to form the backbone of the JH homologs. Due to their low abundance, the substrate specificity of these enzymes has not been studied in enzyme preparations from CA homogenates. The cloning and sequencing of farnesyl diphosphate synthase in A. ipsilon (GenBank Accession no. AJ009962) and B. mori (GenBank Accession no. AB072589) have been reported (Castillo-Gracia and Couillaud, 1999; Kikuchi et al., 2001), and recently Sen and Sperry (2002) have partially purified native farnesyl diphosphate synthase from whole-body preparations of M. sexta. Thus, substrate specificity for this enzyme may be known in the near future. Sen and her colleagues have also examined other enzymes in the JH biosynthetic pathway, including the specificity of prenyltranferase activity of the larval CA of M. sexta (Sen and Ewing, 1997). This enzyme catalyzes the head-to-tail condensation of dimethylallyl diphosphate (DMAPP) and isopentenyl diphosphate (IPP) units to form the skeleton of JH III, and of homoisopentenyl phosphate and ethylmethylallyl diphosphate (EMAPP) to form the skeletons of JH 0, I, II and 4-methyl JH I (Figures 5 and 6). Using homodimethylallyl diphosphate (HDMAPP) and the natural and unnatural homologs of geranyl diphosphate (GPP) (formed from the first coupling of one DMAPP and one IPP in the synthesis of JH III) (Figure 5), they found that this enzyme has a high tolerance for homologous substrates. EMAPP was a better substrate than DMAPP, while the unnatural geranyl homolog was a poorer substrate for the prenyltransferase than the natural homolog, suggesting that this enzyme contributes to the specificity of the JH skeleton. Sen and Garvin (1995) examined the specificity of the farnesol dehydrogenase, an enzyme that

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catalyzes the oxidation of farnesol to farnesal. Their results suggest that this enzyme is a unique dehydrogenase with high specificity for alcohols with D-2,3 unsaturation and alkyl chain hydrophobicity corresponding to at least three isoprenoid units, and hence is a selective enzyme for JH biosynthesis. In a subsequent study, Sperry and Sen (2001) concluded that, in fact, the oxidation of farnesol to farnesal did not involve a unique dehydrogenase, but rather a specific oxygen-dependent enzyme, possibly a flavin- or nonheme iron-dependent oxidase. Further studies on these enzymes will identify targets for compounds with anti-JH activity, as well as foster a better understanding of the key enzymes that are targets of allatotropins and allatostatins (see Section 3.7.7). 3.7.4.2. Possible Feedback Loops in the Biosynthetic Pathways p0135

The formation of farnesyl pyrophosphate is an intermediate step in the synthesis of JH from mevalonate. A mammalian ‘‘orphan’’ nuclear receptor activated by farnesyl pyrophosphate metabolites has been identified (Forman et al., 1995) and termed farnesoid X receptor (FXR) (Weinberger, 1996). FXR is activated by farnesol, farnesal, FA, and methyl farnesoate (Figure 4), and is expressed in isoprenoidogenic tissues. It has been suggested that these farnesyl pyrophosphate metabolites (farnesoids) may be signals for transcriptional feedback control of cholesterol biosynthesis, since cholesterol is synthesized from isoprenoids (Weinberger, 1996). As noted by Gilbert et al. (2000), Weinberger and his colleagues used a transactivation assay in Chinese hamster ovary cells by transfecting with plasmid DNAs expressing FXR, RXR (retinoid X receptor) and a farnesol responsive chloramphenicol acetyltransferase (CAT) reporter to demonstrate that JH, methoprene, and farnesol can all affect mammalian FXR. Farnesoids are JH biosynthetic agonists; they stimulate the formation of JH when added to incubations of CA and are derived from farnesyl pyrophosphate by a metabolic pathway that may serve to excrete surplus isoprenoid precursors (Schooley and Baker, 1985). Since insects cannot synthesize cholesterol de novo and require a dietary source of sterols (Rees, 1985), FXR may be involved in the feedback control of JH biosynthesis via these farnesoids. This possibility certainly deserves further study and opens the door to an exciting field of investigation of the role of orphan nuclear receptors in the regulation of JH biosynthesis (Chawla et al., 2001) (see Section 3.7.7.3)

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3.7.4.3. What Constitutes a Juvenile Hormone Titer?

As discussed by Darrouzet et al. (1998), the identification of new JHs, whether metabolites, conjugates, precursors, or hydroxylated forms, illustrates the variability of compounds released as end products from the CA; these end products are what Darrouzet et al. (1998) term ‘‘bouquets’’ of compounds. The ‘‘bouquets’’ are different in different orders, and undoubtedly vary between species within orders, and even within species by sex and stage in the life cycle (Couillaud, 1995). This is certainly true for M. sexta, where JH I, II, and III and their acids have been measured in larval hemolymph (Baker et al., 1987), and for P. regina, in which the CA secrete JH III, JHB3, and MF (Yin et al., 1995). Thus, it may be naive to think of the circulating titer of a single JH as responsible for the production of a JH effect. Rather, it may be a particular ratio or balance of homologs that is important for biological activity. Sites outside the CA may also contribute to the circulating titers of the different hormones by metabolism, catabolism, hydroxylation, or conjugation. The JH I originally isolated and identified from the abdomens of male H. cecropia pupae (Ro¨ ller et al., 1967) was, in fact, synthesized by male accessory glands from JH acid secreted by the CA (Dahm et al., 1976; Peter et al., 1981). The male accessory glands in the mosquito Aedes aegypti synthesizes JH I, JH III, JHB3, and MF from acetate (Borovsky et al., 1994a), and the mosquito ovary synthesizes JH III from FA (Borovsky et al., 1994b). Non-JHs such as the thyroid hormones, which can be ingested by the insect, may function as a JH in some JH-linked events. Are we safe in assuming that an event is truly JH independent if it occurs in an insect from which the CA have been removed (Davey, 2000a)? Is the JH titer in the hemolymph truly reflective of what elicits a JH-dependent event? Thus, in considering JH titers and any temporally linked JH-dependent event, the event must be viewed from the perspective of a coordinated interplay between products of the CA, the products of other tissues in the body, the catabolism of these products (see Section 3.7.9), and the receptors for these products on the target tissues (see Section 3.7.12.1). The list of products possibly involved in this interplay is long and getting longer.

3.7.5. Measurement of the Juvenile Hormones The lipophilic nature of JH, its occurrence in exceptionally low concentrations in biological samples

p0140

p0145

330 The Juvenile Hormones

(parts per billion or lower), its lability, and its tendencies to bind nonspecifically to substrates, contribute to making this one of the most difficult of all hormones to measure accurately. There are three methods employed to quantitatively assess JH titers: bioassays, physicochemical assays, and radioimmunoassays. Relative JH titers can be determined using the radiochemical assay. 3.7.5.1. Bioassays

The first techniques used to measure JH titers were an assortment of bioassays that quantified a JH effect on a morphological or physiological process (Sla´ ma et al., 1974). Initially, these bioassays were used to assess the activity of transplanted CA, but are now more commonly used to quantify JH indirectly by comparing the biological response of an unknown preparation to a known amount of JH. These types of assays have been well described and critiqued in previous reviews (Feyereisen, 1985; Baker, 1990). Their use is in decline for a number of reasons: (1) they are cumbersome; (2) they are timeconsuming, requiring maintenance of a test insect colony; (3) they are subjective in their scoring; (4) they lack specificity, because they measure only total JH; and (5) they show differential sensitivity to the various homologs. Nevertheless, several assays including the Galleria wax test (Gilbert and Schneiderman, 1961; deWilde et al., 1968) and the Manduca black larval assay (Fain and Riddiford, 1975) are still used, since there are certain instances where bioassays play an important role. These include testing biological extracts for JH activity, especially when the active ingredient is thought to have a non-JH-like structure, or monitoring biological activity of a JH during its purification process. 3.7.5.2. Physicochemical Assays

With growing sophistication in the technology for studying organic compounds, a number of physicochemical methods have been developed for measuring JH in biological samples. Unfortunately, none of these methods is rapid, simple, or inexpensive. Moreover, the literature surrounding the physicochemical detection methods is awash in methods that can be bewildering for a biologist. The current physicochemical methods reflect several decades of experimentation that emphasized specificity and ultrasensitivity at the expense of simplicity. There are normally two phases to the physicochemical assay; the sample extraction/clean-up phase, and the quantification phase. Because JH represents only a minute fraction of the extractable lipids,

especially in whole-body extracts, protocols generally require some type of clean-up procedure(s) such as column chromatography, thin layer chromatography, separatory cartridge purification, HPLC, and/or gas chromatography (GC). Preparation of a biological sample for JH analysis has been discussed in several excellent reviews that cover extraction methods, solvents, and partition techniques (Baker, 1990; Halarnkar and Schooley, 1990). Further cautions with regard to degradation, volatility, and nonspecific adsorption have also been extensively outlined (Granger and Goodman, 1983, 1988; Goodman et al., 1995). Once the clean-up procedure is completed, JH can be quantified by a number of different procedures that employ a gas chromatograph interfaced with an electron-capture detection unit or mass spectrometer (Baker, 1990). In the early analyses of the hormones, GC separation of underivatized JH was coupled with electron impact (EI) or chemical ionization (CI) detection systems to monitor levels of the hormone. While these methods required no derivatization, they often resulted in complex spectra, poor recoveries, and an accumulation of breakdown products (Baker, 1990). Moreover, JH is not particularly stable under most GC conditions, due to instability of the epoxide ring. The use of CI, with its softer ionization process and its improved sensitivity and specificity, yields fewer fragment ions than the harsher EI ionization mode. Nevertheless, problems remain with the misidentification of the JH homologs (Baker, 1990). To enhance sensitivity and specificity, methods were developed that generated JH derivatives with unique chemical tags. The first to be developed were the organohalide derivatives that are detected by a GC interfaced with an electron capture detection (ECD) system. While this method greatly increased sensitivity of the assay, it had serious drawbacks in uniformity of derivatization and in identification of the homologs. To overcome these difficulties, Rembold et al. (1980) and Bergot et al. (1981b) developed GC-mass spectrometry (MS) methods which, while still requiring derivatization of the hormones, were far more efficient and sufficiently selective to allow each JH homolog to be identified using selected ion monitoring. GC-MSEI and GC-MS-CI analyses are now performed with slight variations on these procedures (Neese et al., 2000; Smith et al., 2000; Cole et al., 2002). Teal et al. (2000) have recently developed a method for quantifying JH from biological samples that allows for direct analysis of hormone without derivatization. These investigators use GC interfaced with ion-trap MS and CI. This method

The Juvenile Hormones

requires no clean-up procedure or sample derivatization and is highly sensitive, and thus may be the assay of choice for future studies. While this procedure has been used to titer JH in hemolymph (Burns et al., 2002) and culture medium (Bede et al., 2001), in which lipid content is relatively low, it will be interesting to see if the assay can be successfully used with whole-body extracts. The advantage of the physicochemical assays lies in their sensitivity and their unequivocal identification of the different JHs and metabolites. Their drawback lies in accessibility to the prohibitively costly equipment and the relatively low throughput. 3.7.5.3. Radioimmunoassays

Radioimmunoassays (RIAs) are a rapid, sensitive, and inexpensive alternative to measuring JH, both in biological samples and in medium from incubations of CA. The JH RIA is a competitive protein binding assay in which JH from a biological sample competes with a fixed amount of radiolabeled JH for a limited number of binding sites on JH-directed antibodies. The radiolabeled JH bound to the antibody in the presence of the unknown is compared to a standard curve derived with known amounts of radioinert JH. Since JH itself is not antigenic, antibodies against JH are produced by chemically linking JH to a protein such as human serum albumin or thyroglobulin. The hormone can be conjugated to the protein via the C1 (ester) terminus of JH or through C10,C11 (see Granger and Goodman, 1983, 1988; Baker, 1990; Goodman, 1990, for reviews of the technique). The resulting antiserum is carefully screened for specificity, since it may recognize a particular JH homolog exclusively (Granger et al., 1979, 1982; Goodman, 1990), two or more homologs to differing extents (Granger and Goodman, unpublished data), or all homologs equivalently (Goodman et al., 1993, 1995). Like the physicochemical methods, JH from whole-body or hemolymph extracts must be partially purified prior to conducting the RIA. The partial purification process is essential for a successful assay since a number of lipids can nonspecifically interfere with the assay. Inattention to this process invariably results in excessively large and inaccurate JH titers due to micelle formation of nonspecific lipids that either trap JH or inactivate the JH-directed antibodies. Misidentification of the JH homologs and overestimation of titers can also occur when chromatographic systems contaminated with high levels of JH standards are also used for the purification of biological samples (Baker et al.,

331

1984). It is for these reasons that RIA technology has rightly been criticized (Feyereisen, 1985; Tobe and Stay, 1985). A number of excellent clean-up procedures have been suggested (Strambi, 1981; Strambi et al., 1981; Goodman et al., 1995; Niimi and Sakurai, 1997; Noriega et al., 2001). It is important to note that despite differences in extraction methods and JH-directed antibodies, excellent agreement between assays can be achieved (Goodman et al., 1993). JH RIAs have been used most effectively to measure CA activity in vitro, since the products are secreted into defined incubation medium that lacks the massive amount of lipid found in wholebody or hemolymph extracts. RIAs have been used extensively in studies on CA activity and in control of JH biosynthesis in the tobacco hornworm (Bollenbacher et al., 1987; Granger and Janzen, 1987; Janzen et al., 1991; Granger et al., 1994), locusts (Couillaud et al., 1984; Baehr et al., 1986), honeybees (Huang and Robinson, 1995), and bumblebees (Cnaani et al., 2000). Furthermore, certain extant RIAs recognize JH and JH acid equivalently (Janzen et al., 1991), which can be a benefit in light of the fact that the CA of a number of species secrete JH acid at certain times in their life cycle (Sparagana et al., 1984; Janzen et al., 1991; Ho et al., 1995a, 1995b; Niimi and Sakurai, 1997; Duportets et al., 1998; Audsley et al., 2000). One of the major problems that has plagued research on the chemistry and biology of JH is the lack of commercially available, enantiomerically pure JH; nowhere is this more evident than in RIA technology. Depending upon the antigen used for immunization and immunocompetence of the rabbit, antibodies can be highly specific and capable of recognizing a single enantiomer of each homolog. Thus, using racemic JH as a standard may pose difficulties in establishing an accurate titer. A very complex binding reaction is generated when the radiotracer is racemic, but JH from the biological sample is enantiomerically pure; the analysis becomes even more complex when the radioinert standards are racemic. This problem has been eliminated by the use of a chiral HPLC matrix that can resolve the JH enantiomers (Cusson et al., 1997). Moreover, the use of the appropriate enantiomers leads to increased sensitivity of the assay. Based on the ED50 values (effective dose required to inhibit binding of 50% of the radiolabeled tracer), the naturally occurring enantiomers of JH I, JH II, and JH III in this study were between 30 and 87 times more immunoreactive than the unnatural isomers. In summary, the JH RIA is a relatively rapid, inexpensive, and accurate method for titering JH. If

p0210

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measurement of more than one homolog is required, then confirmation by physicochemical methods is recommended. 3.7.5.4. Radiochemical Assays

The radiochemical assay (RCA), originally developed by Pratt and Tobe (Pratt and Tobe, 1974; Tobe and Pratt, 1974), has been employed for many years to measure JH biosynthesis in vitro by the CA of a wide variety of insects (Feyereisen and Tobe, 1981; Tobe and Feyereisen, 1983; Tobe and Stay, 1985; Yagi and Tobe, 2001). The RCA measures the rate of incorporation of the methyl group from either [14C]methyl methionine or [3H]methyl methionine into JH in isolated CA. The availability of assay components, the relative simplicity of the protocol, its sensitivity (0.1 pmol), and the short incubation time (1–3 h), have made this assay popular among insect endocrinologists. There is a general assumption that isolated glands lacking neural connections function in the same fashion as glands in vivo. This assumption has been challenged by Horseman et al. (1994), who demonstrated that nerve transection in adult L. migratoria led to a rapid, 16-fold increase in JH production. A similar result has also been found in isolated CA of A. aegypti (Noriega, personal communication). Thus, the RCA may, at certain times, seriously overestimate the expected in vivo production of JH (Pratt et al., 1990). Another potential problem is the use of [3H]methyl methionine as the methyl donor (Yagi and Tobe, 2001). The lack of definitive data from the manufacturers on the purity and specific activity of this compound can make determination of the biosynthesized JH difficult. Yagi and Tobe (2001) recommend either comparing incorporation of the [3H]methyl group with the [14C]methyl group in a dual-label experiment or switching altogether to [14C]methyl methionine. It appears that isotopic discrimination by the O-methyl transferase leads to preferential incorporation of the [14C]-labeled methyl group vs. the [3H]-labeled methyl group, but only when concentrations of [3H]methyl methionine above the normal range for the RCA are used. The measurements of JH biosynthesis using the RCA can also be affected by the incubation medium, which in most instances is TC199. TC199 has a high Naþ : Kþ ratio (17.8) and thus is a suitable medium for the CA from insects where the hemolymph ratio of Naþ : Kþ is similar, for example, Leptinotarsa decemlineata or T. molitor (Granger et al., 1986). This medium has been used extensively in the RCA, irrespective of insect

species. For insects such as M. sexta, where the hemolymph Naþ : Kþ ratio is low (0.1) (Nowock and Gilbert, 1976), TC199 has been found to depress JH synthesis in comparison to Grace’s medium, in which the Naþ : Kþ ratio is 0.33 (Granger et al., 1986; Watson et al., 1986). More recently, L-15B, a medium widely used in the culture of arthropod tissues and found to be isoosmotic with cockroach hemolymph, was demonstrated to be superior to TC199 for both long- and short-term cultures of cockroach CA (Holbrook et al., 1997).

3.7.6. Evolution of the Juvenile Hormones 3.7.6.1. Juvenile Hormone Homolog Distribution among the Insect Orders

One or more of the JH homologs have been identified in nearly all species of insects in which they have been sought (Sehnal, 1984) and have even been found in Collembola, an order once thought basal to insects but now considered to represent a more ancestral group, distinct from insects (Nardi et al., 2003). A JH effect on ecomorphosis of a collembolan species, Hypogastrura tulbergi, has been reported (Lauga-Reyrel, 1985), and on this basis, it seems likely that JH or a JH-like molecule functioned in the ancestors of modern insects at least 400 million years ago. Figure 7 illustrates the evolutionary time line in which the various orders of insects were thought to have appeared, the orders in which JHs or JH-like molecules have been identified, and the identity of these compounds. 3.7.6.2. Evolutionary Connections among the Arthropods

Arthropods, which appeared more than 500 million years ago during the Cambrian period, include four extant classes: Hexapoda, Chelicerata, Myriapoda, and Crustacea. Of these, the Hexapoda and Crustacea appear to be the most closely related, based on molecular data, and are now considered together as Pancrustacea (Boore et al., 1995; Friedrich and Tautz, 1995; Tobe and Bendena, 1999). Figure 8 illustrates a time line of the appearance of the extant classes of Arthropoda and those classes in which JH effects have been demonstrated. Outside the class Insecta, JH-like molecules have been identified in Crustacea (see Chapter 3.16), and functions for these molecules have been proposed. Both FA and MF, the immediate precursors of JH III (Figure 4), are major biosynthetic products of the mandibular organs of many crustaceans. The similarities in function and embryological origin of the

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Figure 7 Evolutionary timeline for the appearance of the insect orders based on the earliest documented fossil record (Heming, 2003). Orders in which JH effects have been demonstrated are noted with an asterisk (*). Orders in which JH or JH-like molecules have been conclusively identified are noted by the homolog.

mandibular organs and CA, and the fact that the biosynthetic pathway for MF and FA are the same in both crustaceans and insects (see Section 3.7.4.1), suggests that these products may be the JHs of Crustacea. FA can act as a JH in crustacean reproduction (Laufer et al., 1993), and MF has a juvenilizing role in the prawn (Abdu et al., 1998) and the barnacle (Smith et al., 2000) during metamorphosis from the larval to the juvenile form. MF also delays metamorphosis in larvae of the lobster, Homerus americanus (Borst et al., 1987). Nevertheless, definitive physiological and developmental roles for these two JH precursors remain vague (Homola and Chang, 1997), and further work is needed to determine their exact functions. Although it is not clear whether FA or MF are prohormones, hormones, or simply side products of JH biosynthesis in insects, the esterification of JH acids in the accessory glands of male H. cecropia (Shirk et al., 1983) and the imaginal discs of M. sexta (Sparagana et al., 1985) certainly suggests that FA can be converted to MF, and MF esterified to JH III, in tissues outside the CA. Research is needed on the existence, location, and activity of the enzymes O-methyl transferase and MF epoxidase to elucidate whether FA and MF are prohormones or hormones in their own right.

3.7.6.3. Evolutionary Inferences

FA is released from the nymphal CA of the cockroach, D. punctata, while homo/dihomo MF and FA appear to be products of the CA of adult female and male P. unipuncta (Cusson et al., 1991a). It has recently been shown that in addition to JH III, the embryonic CA of D. punctata produce MF, with a shift to predominantly JH production as development proceeds (Stay et al., 2002) (see Section 3.7.10.2). This evidence has led Stay et al. (2002) to postulate that the production of MF by the early embryonic CA is an ancient trait in Arthropoda, and that the original metamorphic and reproductive hormone in this more basal order was MF rather than JH. In light of the proposal that the ancestral role of JH was involved in embryogenesis (Truman and Riddiford, 1999), MF may have fulfilled this role in the early evolution of the Arthropoda, with a shift to JH III production as metamorphosis in the Insecta evolved. Sehnal et al. (1996) have suggested that the postembryonic development in the ancestors of ametabolous insects occurred in the absence of JH and that the original function of JH in postembryonic life was the regulation of reproduction. As the occurrence of metamorphic molts evolved in ancient

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Figure 8 Evolutionary timeline of the appearance of the extant classes of Arthropoda. Classes in which JH effects have been demonstrated are noted with an asterisk (*). Classes in which JH or JH-like molecules have been identified are noted with a double asterisk (**).

arthropods, it is possible that a hormone (JH) that functioned in the embryo and the mature adult, but played no role in postembryological development, was coopted to regulate development before and after these metamorphic molts (Tobe and Bendena, 1999). In some insect species, JH appears to regulate the types of responses that would facilitate evolution of a larval stage, such as premature differentiation of the embryo and alterations in morphology (Truman and Riddiford, 1999). As outlined below (see Section 3.7.10.7), Truman and Riddiford (1999, 2002) have proposed that an advancement in the time of appearance of JH during embryogenesis may have been key to the evolutionary transformation of a pronymphal stage to the larval stage of holometabolous insects. The production of more complex forms of JH may have occurred with the evolution of holometabolous

development. At the present time, JH III is the only known form of JH in most insect species, particularly the Hemimetabola. With the exception of the HJHs (see Section 3.7.2.2), the more complex forms of JH, JH 0, iso-JH 0, JH I, methyl-JH I, JH III, and JHB3, appear to occur only in two highly derived orders of holometabolous insect, the Lepidoptera and the Diptera. Furthermore, only Diptera appear to have evolved the ability to produce JHB3. The production of the higher homologs and their multiple occurrence in some species appears to be related to the evolutionary development of intricate physiological events and their regulation. The exact physiological or developmental role of the different JHs or JH-like molecules in each class and order needs to be determined to further elucidate the role of JH in arthropod and insect evolution and by extension, the evolution of these molecules themselves.

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3.7.7. Regulation of Juvenile Hormone Biosynthesis A great deal of progress has been made since 1985 in understanding the regulation of JH biosynthesis by neurohormones, neuromodulators, and neurotransmitters (Feyereisen, 1985). At that time it was generally believed, based on a considerable body of in vivo studies, that physiological factors, mainly from the central nervous system, either stimulated (allatotropins) or inhibited (allatostatins) JH synthesis by the CA. Their chemical nature was unknown, but their existence was accepted, based on distinct and predictable changes in the hemolymph JH titer that related directly to physiological events, such as ecdysis of the larva and reproduction in the adult. By 1985, methods for maintenance of the CA in vitro had been established for a number of species, and two approaches to the measurement of JH biosynthesized in vitro had been developed: the radiochemical assay (Pratt and Tobe, 1974) (see Section 3.7.5.4) and the radioimmunoassay (Baehr et al., 1976, 1979; Strambi, 1981) (see Section 3.7.5.3). Using these methods, the JH homologs synthesized by the CA in vitro have been found to be the same as those in hemolymph, although the homolog ratio may vary for CA that synthesize more than one JH. Moreover, developmental changes in JH biosynthetic activity by the CA in vitro parallel changes in the hemolymph JH titers (Feyereisen, 1985; Tobe and Stay, 1985). What factors elicit these changes in JH biosynthetic activity? 3.7.7.1. Neuropeptides

3.7.7.1.1. Allatotropins The first peptide to be isolated that affected JH synthesis in vitro was the allatotropin (AT) of M. sexta (Manse-AT) (Kataoka et al., 1989). While this amidated tridecapeptide did not stimulate JH biosynthesis by larval or pupal CA in vitro, it did stimulate synthesis by adult CA at a concentration of 109 M. The gene for Manse-AT was isolated a decade later and was found to express three different mRNAs that differ from one another by alternative splicing (GenBank Accession no. U62100, U62101, U62102) (Taylor et al., 1996). It has been suggested that the different mRNAs encode three distinct prohormones. Immunocytochemistry with polyclonal antibodies to Manse-AT and in situ hybridization with riboprobes for its mRNAs were used to show that Manse-AT exists in the central nervous system of M. sexta larvae. The former study found immunoreactivity in cerebral neurosecretory cells as well as axons in the corpora cardiaca (CC) and CA (Zˇitnˇan et al., 1995), while the latter found only low levels in

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nonneurosecretory cells of the larval brain and subesophageal ganglion (Bhatt and Horodyski, 1999). Although Manse-AT stimulates the adult CA in this species, its mRNAs were not detected in prepupal, pharate pupal, or adult brains (Bhatt and Horodyski, 1999). Is Manse-AT a functional AT in other lepidopteran species? There is direct evidence from several different studies that this is indeed the case. Manse-AT may be the true allatotropin in P. unipuncta: it has been cloned in this species (GenBank Accession no. AF212926) and ManseAT immunoreactivity has been found in the CA (Truesdell et al., 2000). Manse-AT has also been cloned from the silkworm B. mori (Park et al., 2002) and has been purified from methanolic brain extracts of the fall armyworm, Spodoptera frugiperda, in which it stimulates adult CA (Oeh et al., 2000, 2001). A cDNA encoding 134 amino acids, including an allatotropic peptide, has recently been cloned in this species (GenBank Accession no. AJ488181, AJ508061) (Abdel-Latief et al., 2003), and the mature peptide is identical to Manse-AT. The Spodoptera peptide precursor has 84%, 93%, and 83% amino acid sequence identity with those of M. sexta, P. unipuncta, and B. mori, respectively. Manse-AT also stimulates JH synthesis by both larval and adult CA of L. oleracea (Audsley et al., 1999a, 1999b, 2000), and the adult CA of another moth, Heliothis virescens (Kataoka et al., 1989; Teal, 2002). Manse-AT may be a functional AT in other orders as well. Immunological analyses indicate it is present in the abdominal nervous system of the cockroach Periplaneta americana (Rudwall et al., 2000) and in the brain of the fly P. regina (Tu et al., 2001). Moreover, it stimulates JH synthesis by the larval CA of the honeybee, Apis mellifera (Rachinsky and Feldlaufer, 2000; Rachinsky et al., 2000), and the adult female CA of P. regina (Tu et al., 2002). 3.7.7.1.2. Other roles for Manse-AT Since ManseAT does not affect larval M. sexta CA, could it have another function in the larva? Indeed, Manse-AT has been shown to be a multifunctional molecule in M. sexta, as well as in other species. It exerts an apparently species-specific inhibition of ion transport in the midgut of M. sexta fifth instars (Lee et al., 1998) and accelerates the heart rate in pharate adults, but not larvae (Veenstra et al., 1994). It stimulates foregut contractions in the moths Helicoverpa armigera and L. oleracea (Duve et al., 1999, 2000); acts as a cardioaccelerator in Leucophaea maderae, P. americana (Rudwall et al., 2000),

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and P. unipuncta (Koladich et al., 2002); and is involved in the photic entrainment of the circadian clock of L. maderae (Petri et al., 2002). Thus, a phenomenon observed with a number of other insect neuropeptides is illustrated by Manse-AT: it has multiple functions and these functions are stage-, tissue-, and species-specific. 3.7.7.1.3. Molecular biology of the allatotropins Horodyski and his colleagues have undertaken a detailed study of the molecular biology of the ATs. They have found that while levels of Manse-AT mRNAs are low or nonexistent in the brain, the Manse-AT gene is expressed in a variety of cells in other regions of the central nervous system and frontal ganglion in larval, pupal, and adult stages of M. sexta and that the cellular expression pattern in the nerve cord, as indicated by levels of Manse-AT mRNA, changes during metamorphosis (Bhatt and Horodyski, 1999). Manse-AT-like peptides can be derived from each of three distinct precursor proteins that are the predicted products of alternatively spliced mRNAs (Taylor et al., 1996). The basis for differences in the spliced mRNAs is the inclusion of one or two exons at the same location within the open reading frame. These exons contain a sequence that codes for a peptide with limited structural identity to Manse-AT (Horodyski et al., 2001; Lee and Horodyski, 2002). The three expressed peptides have overlapping but nonidentical activities in JH biosynthesis assays with adult CA in vitro and in the inhibition of active ion transport across larval posterior midgut epithelium in vitro. The presence of the mRNA isoforms resulting from the alternative splicing is regulated in a tissue-and developmentally specific manner, suggesting changing roles for the resulting peptides at different life stages (Lee et al., 2002). Finally, the expression of one of these mRNA isoforms is increased in the ventral nerve cord of M. sexta last instars subjected to treatments that normally reduce feeding and increase levels of JH in the larva: starvation, parasitization, or ingestion of the ecdysteroid agonist RH5992 (K.Y. Lee and Horodyski, 2002). These discoveries are the first to identify the pathway by which these treatments first have their effect. Whether this effect involves downstream regulation of the endocrine system, midgut ion transport, or another target is yet unknown. 3.7.7.1.4. Other allatotropins Until recently, Manse-AT was the only AT identified. However, an allatotropic immunoreactive peptide has been isolated from the abdominal ganglia of the mosquito A. aegypti (Veenstra and Costes, 1999) and found to

have a unique sequence. It has now been shown that Aedae-AT has a stimulatory effect on adult female A. aegypti CA in vitro, with maximum stimulation at dosages of 108–109 M, suggesting that this moiety is the true AT in A. aegypti (Li et al., 2003). Furthermore, while neither the JH precursor FA nor Aedae-AT stimulates JH III synthesis by the CA of newly emerged females, when both are added to the incubation medium, high levels of JH III production occur, indicating that Aedae-AT makes the CA competent to make JH III from FA. There are a number of studies that suggest the existence of at least seven other allatotropins (Stay, 2000; Li et al., 2003). In the wax moth, Galleria mellonella, a protein fraction of larval brains was found to stimulate JH II synthesis by CA in vitro, and monoclonal antibodies raised to that fraction were used to identify a 20 kDa peptide by immunoblotting (Bogus and Scheller, 1996). Two pairs of immunoreactive neurosecretory cells in the G. mellonella brain and in the CC were identified with these monoclonal antibodies. One of the pairs of cerebral neurosecretory cells was immunoreactive with antibodies to Manse-AT as well, but because of the size of the putative Galme-AT, it is unlikely to be a homolog of Manse-AT. The investigators suggest that Manse-AT and Galme-AT have common epitopes due to splicing from the same preprohormone. Recent attempts to isolate the gene for this AT using an expression library and monoclonal antibodies have not been successful (Sehnal, personal communication). A putative AT has been extracted from the subesophageal ganglion of the cricket Gryllus bimaculatus, and stimulates the adult CA of not only Gryllus, but also of another cricket, Acheta domesticus (Lorenz and Hoffmann, 1995). Extracts of the brain–subesophageal ganglion–CC–CA from the true bug, Pyrrhocoris apterus, stimulate JH biosynthesis by the CA of adults raised under long-day photoperiodic conditions (Hodkova et al., 1996), and methanolic extracts of brain or CC from adult vitellogenic females of the locust L. migratoria stimulate JH III release from the CA of vitellogenic females 20–40-fold (Gadot et al., 1987). These extracts also stimulate JH production by the CA of adult male Locusta, but not without some manipulation, since the CA must be chilled at 4  C for 24 h to obtain the stimulatory response (Lehmberg et al., 1992). Surgical manipulations and immunological studies suggest that the brain is the source of this factor (Couillaud et al., 1984; Ulrich, 1985). Allatotropic activity was also extracted from the subesophageal ganglion and CC of adult male M. loreyi, and found to stimulate JH III acid and iso-JH II

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synthesis by male CA in vitro approximately twofold (Kou and Chen, 2000).

JH III bisepoxide synthesis in vitro by CA of another dipteran, the medfly Ceratitis capitata.

3.7.7.1.5. Nonneural allatotropins The lack of effect of Manse-AT and other putative ATs on CA when one would assume an active stimulation of JH biosynthesis implies that there exist ATs of different structures than those isolated to date from neural tissue. Studies suggesting the existence of allatotropic molecules from nonneural sources represent the most intriguing line of research on the control of JH biosynthesis. There currently appear to be two sources for such factors: the ovary and the male accessory sex gland. Ovarian factors that effect JH production have been found in two species, the cockroach D. punctata and the cricket G. bimaculatus. In D. punctata, ovaries stimulate increased JH production in a stage-specific manner: ovaries just prior to ovulation are stimulatory while ovaries of pregnant females are not (Rankin and Stay, 1985). Stimulation is directly related to the size of the basal oocytes; ovaries with basal oocytes smaller than 0.8 mm or larger than 1.5 mm were not stimulatory. More recently, it has been demonstrated that Diploptera ovaries, fat body, and muscle can all stimulate JH synthesis by CA in vitro (Unnithan et al., 1998). Stimulation by the ovary is dose-dependent and sensitive, although the moiety responsible does not appear to be a peptide. In this species, however, the overall control of JH biosynthesis is very much dictated by allatostatins (see Section 3.7.7.2). In G. bimaculatus (Hoffmann et al., 1996), an increase in JH biosynthesis follows the implantation of ovaries into adult males. Whether this effect is mediated by a true AT is not clear, since an extract of ovaries did not stimulate JH synthesis in vitro. A 36 amino acid peptide of known sequence from the male accessory sex gland of D. melanogaster (Chen et al., 1988; Schmidt et al., 1993) stimulates JH synthesis when transferred to the female during mating and also increases JH synthesis in vitro by the CA of adult female D. melanogaster (Moshitzky et al., 1996). The peptide has the same effect in vitro on CA from adult females of the moth H. armigera (Fan et al., 1999). By contrast, fragments of the C-terminal and a truncated N-terminal peptide from this Drosophila sex peptide (SP) inhibit JH synthesis in vitro by CA from both species. Since two N-terminal peptides stimulate JH synthesis, similar to the full-length molecule, it appears that the first five N-terminal amino acid residues are essential for CA stimulation by SP in both species (Fan et al., 2000). Moshitzky et al. (2003) recently reported that the Drosophila SP downregulates

3.7.7.1.6. Future directions of research on allatotropins As proposed by Gilbert et al. (2000), the future directions of research on ATs should concentrate in several areas: (1) the identification of ATs in other orders and the sequences of the Manse-AT-like molecules in those lepidopteran species where Manse-AT has an effect; (2) the exploration of other functions for identified ATs; (3) the elucidation of the interaction of ATs and allatostatins in the overall control of JH biosynthesis; and (4) studies of their receptors and ligand– receptor interactions leading to downstream events that modulate synthesis. To this list should be added research on nonneural ATs. 3.7.7.2. Allatostatins

Allatostatins (ASTs), compounds that inhibit JH biosynthesis by the CA, can be grouped into three families. More than 170 different peptides belong to the largest of the families, the FGLamide ASTs, or the AST-A group. This family is characterized by the presence of a highly conserved pentapeptide in the C-terminus, Y/F-X-F-G-L-amide but displays considerable variability in length and sequence in their N-termini (Tobe and Stay, 2004). These ASTs, first identified by Stay et al. (1991a) in D. punctata, have now been found in many different insect species (Gade et al., 1997; Stay, 2000; Li and Noriega, personal communication). They have also been found in groups outside the insects, including Crustacea (Dircksen et al., 1999; Skiebe, 1999; Duve et al., 2002) and freshwater pulmonate snails (Rudolph and Stay, 1997). In D. punctata alone, there are 14 FGLamide ASTs, with one terminating in an isoleucine (Stay, 2000; Tobe and Stay, 2004). Although Lepidoptera have their own family of ASTs, an FGLamide AST was isolated and identified from M. sexta (Davis et al., 1997). FGLamide immunoreactivity is found in the brain, abdominal ganglia, neurohemal organs, and thoracic motor neurons of M. sexta larvae, but the processes of immunoreactive neurons in the CA are sparse. The authors concluded that in this species, the FGLamides are probably neuromodulatory, myomodulatory, and myotrophic. The second family of ASTs, or AST-B group, has been identified in crickets, locusts, and stick insects and contains a common amino acid sequence of W2W9amide, that is, tryptophan residues at the 2 and 9 positions of the N-terminus (Lorenz et al., 1999, 2000; Tobe and Stay, 2004). Because these peptides in stick insects are variable in length, a

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designation of W-X6-Wamide has been assigned to this family (Tobe and Stay, 2004). At the present time there appear to be 17 of these peptides (Stay, personal communication): five in G. bimaculatus, six in Carausius morosus, with the remainder in orders as diverse as Lepidoptera (M. sexta and B. mori), Diptera (D. melanogaster), Orthoptera (L. migratoria), and Dictyoptera (P. americana) (Stay, 2000). A considerable number of these are not active in the species from which they were isolated, being identified solely on the basis of amino acid sequence. The third family of ASTs, the AST-C group, has been identified in Lepidoptera and has been designated the PISCF family. The first to be discovered, an AST (Manse-AST) in M. sexta, was purified from pharate adult heads by Kramer et al. (1991) and shown to be a 15 residue, nonamidated peptide with a free C-terminus and with no homology to the cockroach ASTs. This neuropeptide is unique in another sense, in that it can completely inhibit JH production in M. sexta. Immunochemical and molecular techniques have shown that Manse-AST is present in other insects, including P. unipuncta (Jansons et al., 1996), L. oleracea (Audsley et al., 1998), Spodoptera littoralis (Audsley et al., 1999a), and D. melanogaster (unpublished information cited in Stay, 2000). As with the other AST families, there is some question as to whether Manse-AST is the functional AST in all these species. Synthetic Manse-AST does not affect JH biosynthesis in vitro by Drosophila ring glands; in P. unipuncta, it does not inhibit larval CA or newly emerged adults, and inhibits adult female CA only 60% at a concentration of 106 M (Jansons et al., 1996). Manse-AST inhibits JH synthesis in vitro by adult CA of Heliothis (Helicoverpa) zea (Kramer et al., 1991) and S. frugiperda (Oeh et al., 1999), but with the latter, only when the CA are first stimulated by Manse-AT. More recently, a synthetic Manse-AST was shown to inhibit JH II synthesis in vitro by CA from larval L. oleracea by approximately 70% (Audsley et al., 2000). Manse-AST has also been demonstrated to modulate JH biosynthesis by the CA of insects of other orders, including A. mellifera (Rachinsky and Feldlaufer, 2000) and P. regina (Tu et al., 2001). Thus, the PISCF ASTs do not appear to be limited to Lepidoptera, and may be the functional AST in other insect orders as well. 3.7.7.2.1. Effect of allatostatins on the juvenile hormone biosynthetic pathway Soon after the discovery of the ASTs, Pratt and his colleagues (Pratt et al., 1989, 1991) showed that addition of exogenous farnesoate or mevalonate reverses the inhibition of

JH biosynthesis in vitro by AST, suggesting that the action of this neuropeptide occurs before the synthesis of the isoprenoid and homoisoprenoid units that condense to form the JH backbone (Granger, 2003) (see Section 3.7.4.1). Tobe and his colleagues (Wang et al., 1994) provided evidence that D. punctata allatostatin 4 (Dippu-AST 4) inhibits the activity of O-methyl transferase, the enzyme catalyzing the penultimate step in JH III biosynthesis in this species (see Section 3.7.4.1), but only at the moderately high concentration of 108 M. The authors concluded that this effect is probably secondary since DippuAST 4 inhibits JH synthesis at a much lower concentration and because a variety of precursors such as FA and (E,E)-farnesol were found in this and other studies to reverse the inhibition by Dippu-AST 4. Wang et al. (1994) concluded that Dippu-AST 4 is acting primarily on the earlier, rather than later, stages of JH biosynthesis. More recent work has confirmed that the most probable targets for the ASTs are the first committed steps in the synthesis of JH III, that is, either the transfer of citrate to the cytoplasm across the mitochondrial membrane or the cleavage of citrate to yield cytoplasmic acetyl CoA, a buildingblock for the isoprenoid and homoisoprenoid units (Sutherland and Feyereisen, 1996). 3.7.7.2.2. Locations of allatostatins in the insect AST-positive cerebral neurosecretory cells have been shown for all three families of ASTs by immunocytochemistry and in situ hybridization. Immunocytochemistry has demonstrated that these cells in M. sexta, D. punctata, and G. bimaculatus are usually located in the lateral protocerebrum, project to the CA on the contralateral side, and arborize in the CC and the CA (see Stay, 2000 for a summary). In some cases, medial neurosecretory cells are immunoreactive, as are some cells in the tritocerebrum, and these cells, together with some of the lateral cells, have been observed to arborize within the brain and thus can be considered interneurons (Stay et al., 1992; Zˇ itnˇ an et al., 1995). With the use of immunocytochemistry, ASTs have been demonstrated in other regions of the central nervous system including the peripheral nerves, the gut, the ovary, and the oviducts (Stay, 2000; Witek and Hoffmann, 2001; Garside et al., 2002) (see Section 3.7.7.2.4). In situ hybridization has been used less frequently to locate ASTs within the insect, usually in combination with immunocytochemistry – for example, the identification of ASTs in hemocytes of D. punctata (Skinner et al., 1997). The wide distribution of the ASTs indicated by these results reinforces their pleiotropic roles.

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3.7.7.2.3. Pleiotropic roles of allatostatins It is clear from the enormous bulk of work on the ASTs that, as with the ATs, their actions are not limited to the modulation of the production of JH and that they frequently do not have an allatostatic role in the insect from which they are isolated. The functions of ASTs have been loosely grouped into six categories: (1) inhibition of JH biosynthesis; (2) modulation of muscle contraction in the heart and gut; (3) modulation of neural activity in the central nervous system; (4) inhibition of vitellogenin synthesis/release; (5) inhibition of the production of ecdysteroids by prothoracic glands and ovaries in vitro; and (6) modulation of digestive enzyme activity (Belle´ s et al., 1999; Stay, 2000; Gade, 2002; Tobe and Stay, 2004). The same AST can exert several different tissue-specific effects at various developmental stages in different insect species. The ubiquitous nature of these peptides in insects and in other invertebrates, plus the conservation of core sequences within them, speaks to their ancient origin. It has been suggested that their function as regulators of JH biosynthesis evolved before the rise of the insects (Tobe and Stay, 2004). The great profusion of ASTs, even within a single species, suggests flexibility and overlap in their functions, although it is clear that in some instances, a single AST or structurally similar group can have a narrowly defined function and only minimal activity in other roles. It has been noted that in insect species more recently evolved than cockroaches, there are fewer ASTs (13–14 in cockroaches compared to 5–9 in Diptera and Lepidoptera), and a concurrent loss of their role as inhibitors of JH biosynthesis (Tobe and Stay, 2004). Since a single AST can have different effects at different times, it is not possible to conclude that the loss of AST function is the direct result of the loss of their numbers, and studies of the molecular evolution of the ASTs are needed to resolve this question (Donly et al., 1993a; Ding et al., 1995; Belle´ s et al., 1999). 3.7.7.2.4. Other types of allatostatins A partially identified factor from the brains of late last instars of M. sexta has been found to inhibit JH production irreversibly in a combination in vivo/in vitro assay (Bhaskaran et al., 1990; Unni et al., 1993). CA exposed in vitro to the brain factor are then implanted into penultimate instars to assay their activity. Because this factor appears to be responsible for stable inhibition of JH biosynthesis until after metamorphosis, it was given the name allatinhibin to distinguish it from ASTs, whose effect is fast and reversible.

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There are also nonneural factors of unknown structure that can inhibit JH synthesis by the CA. The ovary appears to be a prime source for these factors, which is not surprising, given the fact that JH regulates reproduction. Early work by Stay and Rankin (Stay et al., 1980; Rankin and Stay, 1985) indicated that the ovary of D. punctata has a role in the decrease of JH synthesis at the end of vitellogenesis. Allatostatic peptides have been extracted from prechorionogenic ovaries of G. bimaculatus (Hoffmann et al., 1996) and L. migratoria (Ferenz and Aden, 1993). The partially purified Locusta ovarian factor is small (1–1.3 kDa) and inhibits JH synthesis in vitro in a dose-dependent manner. The ovarian extract from G. bimaculatus also inhibits JH synthesis by CA in vitro in a dose-dependent manner, with the potency of inhibition dependent on the stage of the animal from which the CA are taken. Immunoassays using separate polyclonal antisera to Gryllus ASTs A1 (FGLamide) and B1 (W-X6-Wamide) have recently been employed to demonstrate AST-A-like and -B-like immunoreactivity in ovary extracts and partially purified HPLC fractions (Witek and Hoffmann, 2001). The immunoreactive fractions inhibit JH biosynthesis in vitro. The AST- and AST-B-epitopes are immunolocalized to the cortical cytoplasm of oocytes in ovaries of both larval and adult crickets; no neural structures within the oocytes are stained. In D. punctata, where the ovary has a clear role in the decrease of JH biosynthesis at the end of vitellogenesis, the ovarian factor responsible could be either a neuropeptide sequestered by the ovary or an ovarian peptide with an AST epitope. With the use of a competitive reverse transcriptase polymerase chain reaction assay (RT-PCR) to quantify AST expression, it has been demonstrated that both the lateral and common oviducts and ovaries express message for ASTs and that there are changes in the pattern of expression during the reproductive cycle (Garside et al., 2002). Unlike the situation in G. bimaculatus, however, there are no immunoreactive cell bodies in the oviducts or ovary, but there is immunoreactivity in the terminal abdominal ganglion and ventral nerve 7, branches of which innervate the oviducts. Thus, the transcripts quantified by RT-PCR may be generated in the axonal compartment of the allatostatincontaining nerves innervating these structures. 3.7.7.2.5. Molecular biology of the allatostatins The first AST gene to be cloned was that from D. punctata, which encoded for a 41.5 kDa precursor polypeptide (GenBank Accession no. U00444) (Donly et al., 1993a, 1993b). This polypeptide

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contains the 13-member family of AST peptides that exist in this species, with appropriate processing sites for endoproteolytic cleavage and amidation. The ASTs are clustered in the precursor, and separating the clusters are three acidic spacers. These spacers are believed to stabilize the precursor, but they have also been proposed to code for a related peptide (Donly et al., 1993a; Ding et al., 1995). Southern blot analysis revealed that there is a single copy of the AST gene per haploid genome. The prepro-ASTs for all cockroach species examined are similar in size, the organization of the ASTs within them (13 or 14) is conserved, and the separation of groups of peptides by acidic domains is maintained (Ding et al., 1995; Belle´ s et al., 1999). The different AST sequences in the gene of one species were derived from duplication events. Thus, the AST-A family in cockroaches is a good example of a family of peptides with similar sequences derived from a single precursor encoded by a single gene. The Drosophila Genome Project database was screened for sequences corresponding to various insect ASTs. The resulting alignments enabled the cloning of the cDNA for a prepro-Drosophila AST containing four putative A-type ASTs (GenBank Accession no. AF263923) (Lenz et al., 2000). A similar approach was taken to identify a prepro-B-type AST that contains two B-type ASTs (GenBank Accession no. AF312379) (Williamson et al., 2001a, 2001c), and a prepro-C-type AST that contains a single C-type AST with one amino acid residue difference from the M. sexta C-type AST. The sequence of the C-type AST is identical to that of previously identified dipteran ASTs (GenBank Accession no. AF316433) (Williamson et al., 2001b). The mRNA for the G. bimaculatus AST encodes a hormone precursor that contains at least 14 putative hormones (GenBank Accession no. AJ302036) (Meyering-Vos et al., 2001). Five of these were previously identified in this species, while the remaining ones are known from other species. The deduced prepro-AST sequence is similar to that in cockroaches, but shorter than that of locusts. Regions of the acidic spacers that separate clusters of hormones are conserved between cockroaches and crickets, providing additional evidence that the spacers have a real function. Identification of the prepro-AST message has been made in two other species. In the mosquito A. aegypti the message is about 3  103 bases in length and encodes five ASTs (GenBank Accession no. U66841) (Veenstra et al., 1997). The preproAST genes are expressed in both abdominal ganglia and midgut. Three cDNAs of 1506 bases encode

ASTs in the German cockroach, Blattella germanica (Yang et al., 2000) and all have a nucleotide sequence with strong similarity (80%) to that of other cockroach cDNAs for the prepro-AST. 3.7.7.2.6. Control of allatostatin titers The effect of ASTs on JH biosynthesis may occur in a paracrine fashion, i.e., local release within the CA from the axons of cerebral cells that produce these neuropeptides, or in a true endocrine fashion, via the hemolymph. By either means, receptors would mediate the effect of the neuropeptide (see Section 3.7.7.2.7) and it is ultimately the combination of titer and receptor that determines the effect of a neuropeptide. Recent work indicates that degradative mechanisms targeting ASTs can control the levels to which the CA are exposed. Initial work by Bendena et al. (1997) demonstrated that, like most neuropeptides, Diploptera ASTs are susceptible to rapid degradation. One of the ASTs of B. germanica has a half-life of 3–6 min in the intact insect, thus explaining why high doses are required for biological effects in vivo (Peralta et al., 2000). A study of the degradation of Manse-AST by enzymes in foregut extracts of L. oleracea revealed a half-life of 5 min for the AST. Its degradation resulted in two products, both of which are cleaved at the C-terminal side of arginine residues (Audsley et al., 2002). Dippu-AST 7 and Dippu-AST 9 are subject to two primary catabolic cleavage steps: cleavage by a putative endopeptidase, yielding a C-terminal hexapeptide, and subsequent cleavage of this product by an aminopeptidase to yield a C-terminal pentapeptide. Neither of these hemolymph enzymes inactivates the ASTs, since the C-terminal pentapeptide contains the minimal sequence necessary for the inhibition of JH synthesis (Garside et al., 1997a). However, there are membrane-bound enzymes in the brain, gut, and CA that cleave ASTs at the C-terminus, inactivating them in a two-step process (Garside et al., 1997b). Pseudopeptide mimetic analogs of Dippu-ASTs have been synthesized that are resistant to degradation by hemolymph and tissue-bound peptidases, and these can significantly inhibit JH biosynthesis by CA when injected into mated Diploptera females, as measured by an in vitro assay using CA explanted from these females (Nachman et al., 1999; Garside et al., 2000). An outcome of this research is the creation of tools with which the actions of ASTs can be studied in detail, using degradation-resistant analogs. 3.7.7.2.7. Allatostatin receptors The type and number of receptors for ASTs in the CA control

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the timing, duration, and potency these neuropeptides will have on JH biosynthesis. Work on AST receptors has also flourished, particularly with the discovery of such receptors in D. melanogaster, the model insect for genetic and molecular biological investigations. The first study to identify AST receptors was carried out by Cusson et al. (1991b), who used photoaffinity labeling to demonstrate two putative receptors in Diploptera CA. Subsequently, a single receptor for Dippu-AST 7 with a Kd of 7.2  1010 M was found in the CA (Yu et al., 1995), in addition to a single 37 kDa receptor in adult female brains that recognized both Dippu-AST 5 and Dippu-AST 7. Since a single Kd was obtained with AST 5 (9  1010 M), but two with AST 7 (1.5  109 and 3.8  109 M), it was suggested that there were two receptor sites for Dippu-AST 7 in the brain. A subsequent structure– activity study using synthetic analogs of Dippu-AST 2 revealed two receptor types in Diploptera CA (Pratt et al., 1997). The investigators proposed that the C-terminal portion of ASTs contains both the ‘‘message’’ determining its full effect, as well as ‘‘address information’’ determining its binding affinity for the receptor, and that divergent evolution of receptor types occurred with the evolution of multiple ASTs from a common ancestor. More recently, a radioligand-binding assay was used to identify a single class of binding sites for Dippu-AST 7 in midgut membranes with a Kd of 2.1  1010 M (Bowser and Tobe, 2000). In competitive binding assays, Dippu-AST 7 and AST 2 exhibit a higher affinity for the midgut receptor than AST 5, 9, 10, or 11, while the other ASTs did not compete. Clearly, cloning and sequencing of these receptors is needed to sort out the ubiquitous occurrence and varied functions of the multiple ASTs. Two laboratories have cloned G-protein coupled receptors (see Chapter 5.5) from D. melanogaster that are structurally related to mammalian somatostatin/galanin/opioid receptors (GenBank Accession no. AF163775: Birgul et al., 1999; GenBank Accession no. AF253526: Lenz et al., 2000), but which bind a peptide related to the AST family and thus appear to be AST receptors. Lenz et al. (2000) termed their receptor DAR-1. Screening the Drosophila genome database, these investigators found a second G-protein coupled receptor, termed DAR-2, with a 47% amino acid residue similarity with DAR-1 (Lenz et al., 2001). Expression studies of DAR-2 in Chinese hamster ovary cells indicated it is the cognate receptor for four Drosophila A-type ASTs that bind to the receptor differentially. The DAR-2 gene is expressed in embryos, larvae, pupae, and adults, and is mainly expressed in the

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gut of third instars. Since the larval gut of D. melanogaster contains cells expressing the gene for the prepro-ASTs, the authors concluded that DAR-2 mediates AST-induced inhibition of gut motility. In a simultaneous study, both DAR-1 and DAR-2 were expressed in Chinese hamster ovary cells and tested for activation using synthetic Drosophila ASTs and selected Diploptera ASTs (Larsen et al., 2001). Both types of ASTs activate DAR-1 and DAR-2, indicating ligand redundancy and cross-species activity. An apparent A-type AST receptor has been cloned from the cockroach P. americana, with about 60% amino acid similarity in the transmembrane regions to the two identified Drosophila receptors. When functionally expressed in Xenopus oocytes, this receptor exhibits high affinity for cockroach ASTs (GenBank Accession no. AF336364) (Auerswald et al., 2001). An A-type AST receptor has also been cloned from the silkworm B. mori (GenBank Accession no. AH011256) (Secher et al., 2001). This receptor (BAR) shows 60% amino acid residue identity with DAR-1 and 48% with DAR-2. The genomic structure of BAR displays two introns that are coincident with, and have the same intron phasing as, two introns in DAR-1 and DAR-2, and the authors conclude that these three receptors are both structurally and genomically related. BAR mRNA is expressed mainly in the gut, and thus this receptor appears to be a gut peptide hormone receptor. Further studies should focus on identifying AST receptors in the CA to define the second messenger cascade that translates the AST signal in the gland, and to elucidate the evolutionary path by which this large class of peptides evolved to regulate JH biosynthesis. 3.7.7.2.8. Corpora allata sensitivity to allatostatins Humoral factors may play a role in regulating the sensitivity of the CA to ASTs. Unnithan and Feyereisen (1995) found that acquisition of sensitivity to AST by CA from adult mated Diploptera females occurs just before choriogenesis and is dependent on a humoral factor. The investigators could manipulate sensitivity by various experimental techniques, e.g., ovariectomy of vitellogenic females disrupts both JH biosynthesis and acquisition of sensitivity. This line of research deserves more investigation, since it could represent the upregulation of AST receptors by other circulating hormones. 3.7.7.3. Other Factors Regulating Juvenile Hormone Biosynthesis

3.7.7.3.1. Neurotransmitters It has long been thought that regulation of JH biosynthesis is exerted

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via the axons of cerebral neurons that innervate the CA. Yet, it is clear from research on the ASTs that this control could occur via the hemolymph, with release from the CC, the neurohemal organ for cerebral neuropeptides, or other sites of AST-producing cells. Control of JH synthesis by neurotransmitters is also likely, since some of the nerve cells innervating the CA are ordinary neurons (Sedlak, 1985). Octopamine, a primary insect neurotransmitter, was the first neurotransmitter to be identified in the CA of locusts and cockroaches (Evans, 1985). Studies of octopamine effects on JH biosynthesis in vitro have shown either stimulation or inhibition, depending on the species. Octopamine stimulates JH biosynthesis in adult L. migratoria (Lafon-Cazal and Baehr, 1988) and in larvae (Rachinsky, 1994) and adults (Kaatz et al., 1994) of A. mellifera, but inhibits synthesis in adults of the cockroach D. punctata (Thompson et al., 1990) and the cricket G. bimaculatus (Woodring and Hoffmann, 1994). In D. punctata, inhibition by octopamine is bimodal, with peaks occurring at 1010 M and 104 M (Thompson et al., 1990). Recent work on adult females of D. punctata has provided the first indication that JH biosynthesis by the insect CA may be affected by the fast excitatory neurotransmitter l-glutamate, acting via ionotropic receptors to modulate the intracellular Ca2þ concentration (Pszczolkowski et al., 1999) (see Section 3.7.7.4.3). Since high Mg2þ and kynurenate treatments inhibit the l-glutamate-induced stimulation of JH biosynthesis, it was proposed the NMDA (magnesium-sensitive) and non-NMDA (kynurenate-sensitive) receptors are present in the Diploptera CA and are involved in the regulation of JH synthesis. A subsequent study, monitoring rates of JH biosynthesis by Diploptera CA in vitro in response to l-glutamate agonists and antagonists, identified the receptors present as NMDA-, kainate-, and/or quisqualate-sensitive subtypes of ionotropic receptors. These receptors are coupled to calcium ion channels and are thus ionotropic channels rather than metabotropic channels, the latter being coupled to inositol (IP3)-stimulated Ca2þ release (Chiang et al., 2002a). The first suggestion that dopamine might be involved in the regulation of JH production was found in a study of dopamine in the brains of two cockroach species, where its levels fluctuated significantly in relation to events during the JH regulated ovarian cycle (Owen and Foster, 1988; Pastor et al., 1991a). A similar situation has recently been described in the fire ant, Solenopsis invicta (Boulay et al., 2001). Shortly after the study by Pastor et al. (1991a), it was reported that dopamine

affects JH synthesis in vitro by the CA of adult female B. germanica, and that its effect was either stimulatory or inhibitory, depending on the stage of the ovarian cycle (Pastor et al., 1991b). A screen of neurotransmitters in the larval Manduca CA using electrochemical detection-HPLC revealed that dopamine was the only biogenic amine in the gland (Granger et al., 1996). Immunocytochemical analysis further corroborated these studies and demonstrated that dopamine is present in both the brain and CA (Granger, unpublished data). Dopamine affects JH synthesis by larval Manduca CA in vitro differentially, stimulating both JH synthesis and cAMP production by CA in very early fifth stadium glands but inhibiting production of both JH and cAMP after day 2 (Granger et. al., 1996) (Figure 9). Another line of evidence implicating dopamine in the regulation of larval development, possibly through an effect on the CA, derives from studies of the effects of parasitism of the armyworm Pseudaletia separata by the wasp Cotesia kariyai. Parasitism by this species elevates dopamine levels in the nerve cord and hemolymph, slows normal development, and delays pupation of the host (Noguchi et al., 1995; Noguchi and Hayakawa, 1996). Dopamine levels are significantly elevated in the hemolymph of M. sexta larvae parasitized by the wasp Cotesia congregata (Hopkins et al., 1998), although no developmental effects are related to the increase. Significantly higher levels of dopamine have also been found in the hemolymph and central nervous system of diapause-bound pupae of the armyworm Mamestra brassicae (Noguchi and Hayakawa, 1997) (see Chapter 3.12). The relationship of elevated dopamine levels and delayed pupation is unclear, although persistent levels of JH are known to delay pupation in Lepidoptera. It is tempting to think that elevated levels of dopamine are stimulating CA activity. However, the converse could equally be true: persistent JH synthesis delaying pupation could elevate dopamine levels. A similar situation occurs in the honeybee: JH has been implicated in the control of honeybee division of labor, where elevated levels of octopamine and serotonin in the antennal lobes of adult workers are associated with foraging behavior (Schulz and Robinson, 2001) (see Chapter 3.13). Treatment of honeybees with methoprene elevates octopamine levels, and these authors concluded that JH modulates octopamine levels in the honeybee brain, and that octopamine acts downstream of JH to influence behavior (Schulz et al., 2002). Neurotransmitter receptors exist in the CA, which is not surprising given that these glands

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Figure 9 Effect of dopamine synthesis on JH biosynthesis in vitro. Manduca sexta CA from different days during the last larval stadium were incubated with increasing concentrations of dopamine. JH III/JH III acid synthesis was measured by radioimmunoassay (RIA) and expressed as ng of JH III radioimmunoassay equivalents. (Adapted from Granger, N.A., Sturgis, S.L., Ebersohl, R., Geng, C.X., Sparks, T.C., 1996. Dopaminergic control of corpora allata activity in the larval tobacco hornworm, Manduca sexta. Arch. Insect Biochem. Physiol. 32, 449–466.)

are extensively innervated by the axons of cerebral neurons and neurosecretory cells. The receptors for biogenic amines belong to a large superfamily of G-protein coupled receptors, and the nucleotide sequences of the octopamine, tyramine, dopamine D1 and serotonin receptors are known for D. melanogaster, A. mellifera, H. virescens, B. mori, and L. migratoria (von Nickisch-Rosenegk et al., 1996; Blenau and Baumann, 2001). Binding of dopamine to D1 receptors increases adenylyl cyclase activity, while binding to D2 receptors either has no effect or inhibits adenylyl cyclase (Gingrich and Caron, 1993; Vernier et al., 1995; Huang et al., 2001). D2-like receptors have not yet been cloned in any insect, although pharmacological studies suggest they exist (Blenau and Baumann, 2001), and receptors for biogenic amines in the CA have not been pharmacologically characterized except for the dopamine receptors in the CA of M. sexta last instars (Granger et al., 2000). Because dopamine both stimulates and inhibits JH synthesis and adenylyl cylase activity, depending on developmental stage, these glands undoubtedly possess both D1-like and D2-like receptors (Granger et al., 1996). The CA D2-like receptor was found to have some pharmacological resemblance to vertebrate D2 receptors, but certain vertebrate D1 receptor agonists/ antagonists were found to be equally effective as D2 receptor agonists/antagonists. By contrast, the CA D1-like receptor was found to be pharmacologically distinct from vertebrate D1 receptors (Granger et al., 2000). This fits with the overall picture of dopamine receptors that have been cloned from insects–they display pharmacological properties that

set them apart from vertebrate receptors (Blenau and Baumann, 2001). 3.7.7.3.2. Interendocrine control by ecdysteroids If, according to the axiom of Carroll Williams (1976), JH is the handmaiden of ecdysone, then one might ask: Do ecdysteroids regulate JH biosynthesis? The evidence that ecdysteroids may regulate CA activity comes primarily from Granger, Bollenbacher, and their colleagues, who determined that regulation is interendocrine and occurs via the brain. The increase in the ecdysteroid titer, specifically 20-hydroxyecdysone (20E), that elicits pupal commitment in the last stadium of M. sexta and other Lepidoptera, also stimulates the synthesis of JH acids by Manduca CA. This effect, exerted only when the CA are incubated as a complex with the brain–CC, occurs in response to physiological concentrations of 20E, and results in the postcommitment increase in the hemolymph titers of JH acids that have been shown to be necessary for development to the pupa (Whisenton et al., 1985; Watson et al., 1986; Granger et al., 1987). A subsequent investigation of the kinetics of the 20E effect revealed that maximum stimulation was achieved in 1 h, supporting the idea of an indirect effect, probably via the brain, and suggesting the existence of a mediating factor (Whisenton et al., 1987). Physiological concentrations of 20E were also shown to stimulate JH synthesis in Manduca fourth stadium CA, taken before the normal rise in CA activity at the end of the stadium. This suggests that interendocrine control of JH biosynthesis during larval molting is also exerted by 20E (Whisenton et al., 1987), and that this effect

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is specific for a biologically active ecdysteroid, since two biologically inactive ecdysteroids, 22-isoecdysone and 5-a-ecdysone, failed to stimulate the CA. The effective concentrations that stimulate the CA differ by about 200-fold, 1 mg ml1 during larval development and 0.02 mg ml1 during pupal commitment, in keeping with the hemolymph concentrations of 20E at those times (Bollenbacher et al., 1981). Interestingly, both of these concentrations stimulate CA from last instars at the time of commitment, and there are no other maxima between these two concentrations (Figure 10) (Granger, unpublished data). Given the fact that Manse-AT does not affect Manduca larval CA, it would appear that 20E acts in lieu of an AT in larvae. Incubation of brain–CC–CA complexes from pharate Manduca pupae with physiological concentrations of 20E resulted in a 40% decrease in the production of JH acids (Granger et al., 1987). This decrease did not occur if the glands were incubated with, but separated from, the brain–CC complex. While the level of inhibition seen in these experiments is not sufficient to account for the inactivation of the CA by the end of the stadium, this mechanism could act in concert with Manse-AST, which does inhibit larval glands. Granger et al. (1996) have demonstrated that the dopamine stimulation of JH synthesis by CA from day 0 last instars of M. sexta may be downregulated by 20E. Ecdysteroid receptors (see Chapter 3.5) have been detected in the nuclei of Manduca CA as early as day 3 of the last stadium (Bidmon et al., 1992),

Figure 10 Effect of increasing concentrations of 20-hydroxyecdysone on JH I acid biosynthesis in vitro by CA incubated as a brain–CC–CA complex, from gate II, day 3 fifth instars of M. sexta. Each datum point represents the mean (
) SEM, where n = 5–6 incubations. Results are expressed as ng JH I RIA equivalents synthesized per complex in a 6 h incubation. (From Granger, unpublished data.)

suggesting the possibility that the negative effect of ecdysteroids on JH biosynthesis involves nuclear receptors and transcriptional control. In a study of the ecdysteroid receptors in Manduca CA, it was found that both components of the heterodimeric receptor complex – ecdysteroid receptor (ECR) and Ultraspiracle protein (USP) – are expressed (Gilbert et al., 2000; Granger and Rybczcynski, unpublished data). The CA contain two or more USP isoforms that are probably the phosphorylation states of two primary translation products. Preliminary results show that the p49 USP isoform in the CA increases during the feeding period and after commitment to pupation in the last larval stadium (Figure 11). The larger USP isoforms show a maxima surrounding the time of the commitment peak in ecdysteroid titer and during the premolt ecdysteroid peak. This pattern directly contrasts with that of the ECRs which show few differences in relative abundance or isoform diversity in this stadium (Gilbert et al., 2000). Culture in vitro of day 1 CA with ecdysteroids causes a decrease in the predominance of the larger USP isoforms and an increase in p49 (Figure 11). This response is also seen in prothoracic glands (Song and Gilbert, 1998). It may be that the changes in ecdysteroid titer are responsible for changes in USP isoform number and abundance during the last stadium and thus might modulate the number and/or kind of genes (for dopamine receptors, for example, which are affected by the ecdysteroid titer) activated by a given concentration of ecdysteroid receptor. With regard to other possible molecular interactions of ecdysteroids and the JH biosynthetic pathway, the evidence is largely conjectural. As discussed above (see Section 3.7.4.2), a mammalian farnesoid receptor, FXR, is affected by JH, methoprene, and farnesol in a mammalian transactivation assay. Weinberger (1996) has demonstrated that bile acids, 3-a-hydroxysteroids, and oxysterols are endogenous FXR affectors. Is it possible that another class of steroids, the ecdysteroids, also affect FXR, and could this be a mechanism by which ecdysteroids affect JH biosynthesis? 3.7.7.3.3. Regulation of juvenile hormone biosynthesis by juvenile hormone There is a paucity of direct evidence that JH regulates its own synthesis. An early study by Granger et al. (1986) indicated that neither exogenous JH I nor JH III can inhibit JH I acid biosynthesis by CA from last instars of M. sexta. A later, more detailed study of JH feedback in this system confirmed this result (Granger, unpublished data). Richard and Gilbert (1991), using Drosophila ring glands, demonstrated that

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Figure 11 Ultraspiracle expression in the corpus allatum during the fifth larval stadium of M. sexta. The proportion of the p49 isoform increases as the stadium progresses, and this shift can be replicated in vitro by incubating CA with 20-hydroxyecdysone as described by Song and Gilbert (1998). CA were dissected on the indicated days and subjected to sodium dodeylsulfate polyacrylamide gel electrophoresis (SDS-PAGE) followed by immunoblotting with an anti-USP antibody (Song and Gilbert, 1998). V0–V9, days of the fifth larval stadium; C, control, V1 CA; E, V1 CA incubated with 10 mM 20E for 18 h. Arrows indicate isoforms of USP. (Rybczynski, Moshitzky, and Granger, unpublished data.)

JH III and JHB3, but not MF, reversibly inhibited biosynthesis of JH III, JHB3, and MF. These results are in agreement with the generally accepted mechanism of negative feedback regulation of an endocrine gland by its product(s) (see Chapter 3.8). As discussed above (see Section 3.7.4.2), mammalian orphan nuclear receptors now appear to be involved in the coordination of the lipid-based metabolic signaling cascade (Chawla et al., 2001). One of these nuclear receptors, RXR, is affected by farnesol and JH, and thus might be of significance in considering regulatory feedback loops of JH on JH biosynthesis. Molecular approaches have led to spectacular advances in our understanding of insect endocrinology, and the future holds great promise with respect to the elucidation of interendocrine control mechanisms. 3.7.7.4. Second Messengers

Receptors for neuropeptides and neurotransmitters are usually coupled to intracellular reaction cascades, regulating the level of second messengers that transduce these signals (see Chapter 3.2). The influence of a variety of second messenger systems on JH biosynthesis has been investigated both independently and as signal transducers, including cyclic AMP (cAMP), cyclic GMP (cGMP), phospholipid/ protein kinase C, inositol phosphates, diacyl glycerol (DAG), and calcium (Ca2þ).

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3.7.7.4.1. Cyclic AMP The role of cAMP in the regulation of JH biosynthesis has been studied most extensively in the cockroach D. punctata. In an initial study, compounds that increased intracellular cAMP were found to inhibit JH synthesis in vitro (Meller et al., 1985). These data suggest that the effect of the ASTs is mediated by cAMP. Of various compounds tested, 8-bromo-cAMP, which is resistant to hydrolysis by phosphodiesterases, and forskolin, a diterpene that activates adenylyl cyclases, were both potent inhibitors of JH synthesis in vitro. Furthermore, developmental changes in CA sensitivity to forskolin were found to parallel those to ASTs (Stay et al., 1991a), and exposure of CA to crude brain extracts resulted in both an increase in cAMP

and a decrease in JH production (Aucoin et al., 1987). However, a subsequent study by Tobe (1990) showed that cAMP levels were low in virgin female CA, whose JH biosynthesis is inhibited. More cGMP than cAMP was found in these CA, and cGMP was lowest when JH biosynthesis was at a maximum in mated females. Levels of cAMP and cGMP were then measured in the CA of both virgin and mated females exposed to synthetic DippuASTs 1–4 (Cusson et al., 1992). No significant changes in the levels of either cyclic nucleotide were found, strongly suggesting that neither of these cyclic nucleotides is involved in the signal transduction of the ASTs. Octopamine inhibition of JH biosynthesis in this species is bimodal (see Section 3.7.7.3.1), but only the inhibition in response to the higher concentration of octopamine (104 M) results in a rise in the levels of intracellular cAMP (Thompson et al., 1990). Thus, the caveat expressed by Meller et al. (1985), that the effects of pharmacological agents might only mimic the cellular response to inhibitory factors, appears to hold in D. punctata, where only a moderately high dose of octopamine exerts an inhibitory effect via an octopamine-sensitive cAMP. By contrast, cAMP has a stimulatory effect on larval CA activity in M. sexta and is involved in transduction of the dopamine signal. Granger et al. (1994) found that relatively inactive CA, such as those of day 4 of the last stadium or from the black larval (bl) mutant, were sensitive to a variety of compounds in vitro that elicited, or mimicked the effects of elevated intracellular cAMP levels. It was also discovered that the stage-specific effects of dopamine on JH/JH acid biosynthesis by M. sexta larval CA in vitro were mediated by a calcium/calmodulin sensitive adenylyl cyclase (Granger et al., 1995b). Dopamine at concentrations of 106 and 107 M stimulated adenylyl cyclase activity in homogenates of glands taken from day 0 last instars, when this neurotransmitter stimulates JH biosynthesis, but inhibited enzyme activity in homogenates of day 6 glands, when dopamine inhibits CA activity (Granger et al., 1995b, 1996). A similar approach demonstrated that octopamine had no effect on

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enzyme activity in gland homogenates over a wide range of concentrations. It was subsequently found that effects of dopamine receptor agonists and antagonists on JH biosynthesis in vitro were mirrored by changes in adenylyl cyclase activity (Granger et al., 2000). Thus, in this species, cAMP appears to be important in neurotransmitter effects on the CA. Transduction of a neurotransmitter signal appears to be mediated by cAMP in the CA of adult male L. migratoria as well (Lafon-Cazal and Baehr, 1988). In this species, stimulation of JH biosynthesis by dopamine correlates with an increase in adenylyl cyclase activity. In honeybee larvae, both octopamine and serotonin elicit a dose-dependent increase in JH release from intraglandular stores of JH in the CA (Rachinsky, 1994). Incubation of these glands with the phosphodiesterase inhibitor 3-isobutyl-1-methylxanthine (IBMX) and a threshold concentration of octopamine (108 M) potentiated the octopamine effect on JH release, indicating that the action of octopamine is mediated by cAMP. 3.7.7.4.2. Inositol phosphates and diacylglycerol Other possible candidates for the transduction of neuropeptide signals in insects are IP3 and diacylglycerol (DAG), which are products of phosphatidylinositol-4,5 bisphosphate hydrolysis. The binding of the neuropeptide to a cell membrane receptor, which then interacts with membranebound G proteins, initiates this cascade. Increased IP3 then elicits the release of Ca2þ from intracellular storage sites, while DAG activates protein kinase C (PKC) (Berridge, 1993). The IP3 second messenger pathway exists in insects (Berridge, 1993). Work by Feyereisen and Farnsworth (1987) demonstrated that phorbol esters, which activate PKC, are potent inhibitors of JH biosynthesis in D. punctata, suggesting this pathway as central to the transduction of the AST signal. In a subsequent study, Rachinsky et al. (1994) found that treatment of Diploptera CA in vitro with inhibitors of DAG kinase elevates the concentration of DAG, resulting in a significant, dose-dependent inhibition of JH biosynthesis. A further enhancement of this effect was obtained when the CA were exposed in vitro to both DAG kinase inhibitors and ASTs, suggesting that the DAG pathway is indeed involved in the cellular response to the AST signal. Treatment of the CA with thapsigargin, a drug that mobilizes calcium stores without generation of inositol phosphates, significantly stimulates JH biosynthesis by, and also reverses the effect of ASTs on, highly active CA from mated females. It does not affect inactive CA from virgin females. The authors concluded

that IP3, the other product of phosphoinositide hydrolysis, could modulate JH biosynthesis at specific development time points and thus could be responsible for the decreasing sensitivity of the CA of mated females to ASTs (Stay et al., 1991b). Furthermore, the stage-specific effects of IP3 on intracellular free Ca2þ in turn modulate the antagonistic action of DAG (Rachinsky and Tobe, 1996). In adult male and female M. sexta, Manse-AT induces the production of inositol phosphates, including IP3, in CA in vitro, and two intracellular Ca2þ releasing agents were found to stimulate JH biosynthesis in vitro, similar to the effect of Manse-AT (Reagan et al., 1992). On this basis, these investigators concluded that Manse-AT activates the IP3 pathway in adult Manduca CA, increasing the intracellular concentration of free Ca2þ and thus stimulating JH biosynthesis. Staurosporine, a PKC inhibitor, had no effect on the inositol phosphates, but did stimulate the rate of JH biosynthesis in vitro and indicated a role for DAG as well. DAG involvement must be confirmed and the mechanism of staurosporine-induced stimulation must be established. 3.7.7.4.3. Calcium Calcium is unique as a second messenger in the CA, in that a transmembrane channel is involved, which may open either in response to ligand–receptor binding or to a change in membrane potential. The resulting increase in cytosolic free Ca2þ then interacts with the same cascades of biochemical events as cyclic nucleotides and inositol phosphates. While JH biosynthesis by Diploptera CA in vitro can be stimulated by the release of intracellular Ca2þ stores (Rachinsky et al., 1994; Rachinsky and Tobe, 1996), it appears that calcium stimulation of JH production occurs primarily by its influx from the extracellular environment. Calcium was first indicated as a player in the control of JH biosynthesis in D. punctata with the observation that elevated extracellular Ca2þ substantially reduces the effects of brain extract (ASTs) on JH biosynthesis (Aucoin et al., 1987). It was then shown that JH synthesis is nearly totally inhibited in Ca2þ-free medium but is stimulated in media with increasing concentrations of this ion (Kikukawa et al., 1987). Studies by Thompson and Tobe (1986) indicated that Ca2þ acts as a second messenger to stimulate JH biosynthesis by entering CA cells through voltage-gated Ca2þ channels. Recent work by Chiang and colleagues demonstrated that another type of Ca2þ-permeable membrane channel is involved in regulation of JH production by Ca2þ; the receptor-operated, ionotropic l-glutamate receptor (Pszczolkowski et al., 1999; Chiang et al., 2002b).

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Elevated Ca2þ concentrations in incubation medium stimulate the synthesis of JH in vitro by the ring glands of D. melanogaster (Richard et al., 1990) and the larval blowfly Lucilia cuprina (East et al., 1997), the larval CA of M. sexta (Allen et al., 1992a), and adult male CA of the cricket G. bimaculatus (Klein et al., 1993). By contrast, JH biosynthesis by the CA of L. migratoria appears to be relatively independent of changes in extracellular Ca2þ, although calcium ionophores can elicit and increase synthesis, depending on the physiological status of the adult source (gregarious but not solitary) (Dale and Tobe, 1988a). With M. sexta CA, Ca2þ levels may vary with the stage of development. Larval CA were found to require extracellular Ca2þ 0.1 mM for maximal JH biosynthesis in vitro, while JH acid synthesis by glands taken after pupal commitment continues in the absence of extracellular Ca2þ (Allen et al., 1992a). Both calcium ionophores and caffeine, which initiate the release of Ca2þ from intracellular stores, stimulate JH acid synthesis by postcommitment CA, suggesting that intracellular Ca2þ may be the principal source of this ion after commitment. Calcium channel antagonists and calcium channel blockers decrease JH biosynthesis by both larval and postcommitment CA, indicating that Ca2þ channels exist in the CA cell membrane and that these channels may be both voltage-gated and receptor-operated. Measurements of cytosolic free Ca2þ in isolated Manduca CA cells by use of the fluorescent Ca2þ indicator Fura-2 and digitized microscopy revealed that the highest levels of intracellular Ca2þ occur on days 0 and 6 of the last stadium, when in vitro synthesis of JH and JH acid is at the highest levels (Allen et al., 1992a, 1992b) (Figure 12). Concentrations of free Ca2þ in Manduca hemolymph peak on days 0 and 7 of the last stadium (Allen et al., 1992a); thus, there is an optimal concentration of Ca2þ in the hemolymph at the time when the CA require extracellular Ca2þ for maximal synthesis. This finding suggests that extracellular free Ca2þ could be responsible, at least in part, for the high activity of the gland at this time. Furthermore, free cytosolic Ca2þ levels decrease significantly by day 1, when the hemolymph titers in JH have dropped precipitously (Baker et al., 1987). However, the concentrations of free Ca2þ in the hemolymph throughout the last stadium never drop below millimolar concentrations (Allen et al., 1992a). Thus, it is clear that free Ca2þ is a significant player in the control of CA biosynthetic activity, but probably as a partner to another signal. It is interesting that steroid hormones are known to stimulate Ca2þ influx in certain

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biological systems (Blackmore et al., 1990) and that free Ca2þ is highest in the CA cells at times when the hemolymph ecdysteroid titer in M. sexta is high (Bollenbacher, 1988). Thus, another possible mechanism for interendocrine control of JH synthesis by ecdysteroids (see Section 3.7.7.3.2) could involve the calcium ion. A similar approach to measuring free Ca2þ in isolated cells of Diploptera CA was used to study the effects of thapsigargin and Dippu-AST (Rachinsky and Tobe, 1996). It was found that thapsigargin elevated intracellular Ca2þ, confirming the stimulatory effect of thapsigargin on JH biosynthesis via elevated Ca2þ; this effect could not be blocked by the later addition of Dippu-AST. The simultaneous addition of Dippu-AST with thapsigargin prevented the thapsigargin-induced increase in intracellular Ca2þ, although, as mentioned previously, the inhibition of JH biosynthesis by Dippu-AST is abolished by thapsigargin (Rachinsky et al., 1994). These conflicting results are not yet resolved. It has been known for many years that the calcium ion also can act indirectly as a second messenger, by binding to calmodulin (CaM) and eliciting a conformational modification of CaM (see Chapter 3.2). This enhances the affinity of the Ca2þ/CaM complex for target effectors, such as membraneassociated and cytosolic enzymes and transmembrane ion channels, including that for Ca2þ (Stoclet et al., 1987). It is of interest to note that a Ca2þ/ CaM-sensitive adenylyl cyclase has been identified in the CA of Manduca last instars (Granger et al., 1995b). 3.7.7.4.4. Potassium Postassium also affects JH biosynthesis. Treatment of adult female L. migratoria CA in vitro with high levels of Kþ stimulates JH biosynthesis, either by stimulating the release of ASTs from nerve endings within the CA or by a direct effect on the CA cells (Dale and Tobe, 1988a). A Kþ-elicited inhibition of JH synthesis was also reported for D. punctata (Rankin et al., 1986), but in both cases, the effect of high concentrations of Kþ proved to be Ca2þ-dependent because it was abolished by Ca2þ channel blockers (Dale and Tobe, 1988b). In summary, based on the evidence to date, it appears that the allatotropic and allatostatic neurotransmitters, which are in evolutionary terms the more primitive intercellular signaling molecules, operate via cyclic nucleotide second messengers. The neuropeptidergic signals that evolved later are transduced by phosphoinositides and DAG. Calcium plays a central role in the function of both types

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Figure 12 Measurements of cytosolic free Ca2+ in isolated Manduca CA cells. Pseudocolored ratio images of CA cells from fifth stadium larvae and early pupae. Cells loaded with Fura-2 and incubated in Grace’s medium containing 0.1 mmol l1 Ca2þ. (a) Day 0; (b) day 1; (c) day 2; (d) day 4; (e) day 6; (f) day 0 pupal period. The color spectrum in A corresponds to the concentration range (nmol l1) of Ca2+. Scale bars represent 120 mm. (Reproduced with permission from Allen, C.U., Janzen, W.P., Granger, N.A., 1992a. Manipulation of intracellular calcium affects in vitro juvenile hormone synthesis by larval corpora allata of Manduca sexta. Mol. Cell. Endocrinol. 84, 227–241.)

of second messengers, and may itself contribute to the upregulation of gland activity at certain critical times when JH biosynthesis levels are high. 3.7.7.5. Putting It All Together

Given the long list of factors that can affect the biosynthetic activity of the CA, and perhaps also

its product(s) – something that has not yet been explored, attempts to create an overall mechanism of endocrine control is fraught with hazards. An excellent summary of the roles of calcium and phosphoinositides in second messenger signaling in the regulation of JH production was made by Rachinsky and Tobe (1996), using three different

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Figure 13 Hypothetical scheme for the regulation of JH biosynthesis by the CA. Factors thought to be involved include neurohormones, neurotransmitters, ecdysteroids, and second messengers. L, ligand; R, receptor; ROC, receptor-operated channel; VGC, voltage-gated channel; c, catalytic subunit; R1,2, regulatory subunit; C, cyclase; Gs, stimulatory G protein; Gi, inhibitory G protein; Gq, G protein associated with phospholipase C; PKC, protein kinase C; DAG, diacylglycerol; IP3, inositol triphosphate; PIP2, phosphatidylinositol-4,5 bisphosphate; TK, tyrosine kinase; PLC, phospholipase C.

insects as model systems: L. migratoria, in which JH biosynthesis appears to be governed principally by allatotropic signals; D. punctata, where this role is filled by ASTs; and M. sexta, in which both types of signals occur, perhaps as the result of the complex nature of the products of the CA in this species and possibly coupled with its holometabolous development. They concluded that differences in signal transduction between species reflect, in part, differences in the signals themselves, and that these differences may be stage-specific. With the complicated nature of this discussion, which excludes a consideration of signals other than neuropeptides, neurotransmitters, and selected hormones, we have chosen to create a scheme for a CA cell that is representative of all insects, responding to these various types of signals and containing all of the second messenger systems shown to exist thus far (Figure 13). Thus, this cell responds to ATs and ASTs, neurotransmitters, ecdysteroids, and Ca2þ, operating via either voltage-gated or receptor-operated channels. It contains adenylyl cyclase, phosphoinositide/DAG, and Ca2þ second messenger

systems, involving PKC. The complexity of this scheme indicates how much work is yet to be done on the basic mechanisms regulating JH biosynthesis at the level of the gland.

3.7.8. Hemolymph Transport Proteins for the Juvenile Hormones The physicochemical nature of JH presents the insect with a serious problem, that of dispersing a lipid hormone via an aqueous circulatory system. While the JH homologs are water-soluble at levels that far exceed the physiological titers normally encountered (Kramer et al., 1976), their amphiphilic nature promotes surface binding (Law, 1980). When dispersed in an aqueous medium, JH displays a marked propensity for nonspecific binding to nearly any surface, making its distribution problematic. Early in the evolution of insects and crustaceans, hemolymph proteins arose that interacted with JH in a noncovalent fashion, yet allowed them to be dispersed in the aqueous environment of the hemolymph (Li and Borst, 1991; King et al., 1995). These

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transport proteins, with their hitch-hiking hormones buried in a hydrophobic pocket, provided early arthropods with a readily available source of hormone that could dissociate from the protein and interact with target tissues. These transport molecules have evolved into highly specific hormone carriers that have been extensively studied since their discovery in the 1970s (Trautmann, 1972; Whitmore and Gilbert, 1972). The hemolymph of most insect species contains two classes of JH-binding macromolecules: (1) nonspecific binding molecules characterized by a high equilibrium dissociation constant (Kd ¼ >106 M; low affinity), and (2) specific binding molecules exhibiting a low equilibrium dissociation constant (Kd ¼ JH I > JH III acid > JH diol. Curiously, no displacement is observed with JHB3 even though it is the predominant in vitro product of the blowfly CA (Trowell et al., 1994). The dissociation constant for JH–JHBL is approximately 30 nM and, as with the other JHBLs studied to date, the fat body is the site of synthesis. s0255 p0595

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3.7.8.1.2. Larval hexamerins as juvenile hormone transporters A second type of high affinity, high molecular weight JH transport molecule has been discovered, having the characteristics of a hexamerin. The hexamerins, composed of six 70–80 kDa subunits, are widely distributed throughout the phylum Arthropoda and have been found in insects, crustaceans, and certain chelicerates (Burmester, 2002); however, they are not typically employed as hemolymph JH transporters. To date, only species in the order Orthoptera, including L. migratoria (Koopmanschap and deKort, 1988; Braun and Wyatt, 1996) and Melanoplus sanguinipes (Ismail and Gillott, 1995), are known to exploit hexamerins as JH transport proteins. The most thoroughly studied of the JH-binding hexamerins (JHBHs) is that of L. migratoria. This protein, a 74.4 kDa protein, as deduced by cDNA analysis (Braun and Wyatt, 1996), contains 15% lipid (Koopmanschap and deKort, 1988). Binding analysis indicates the dissociation constant for (10R)-JH III–JHBH is 1–4 nM. According to Koopmanschap and deKort (1988), the hexamerin is present at relatively low concentrations that never exceed 2% of the total hemolymph protein, yet its hexameric structure allows a single native molecule to bind up to six molecules of hormone. As a result, this JHBH contains very large number of unoccupied hormone binding sites. The 4.3 kb hexamerin mRNA encodes a 668 amino acid protein and contains 2 kb of 30 untranslated region (GenBank Accession no. U74469) (Braun and Wyatt, 1996). Functional assays to locate the JH binding site suggest that it resides in the N-terminus, since a truncated JHBH lacking this region does not bind the hormone. A comparison of its amino acid sequence with other members of the hexamerin superfamily indicates that the JHBH from L. migratoria represents a new form that is most closely aligned with the hemocyanins (Braun and Wyatt, 1996).

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3.7.8.2. High Affinity, Low Molecular Weight Hemolymph Juvenile Hormone Binding Proteins

While the high affinity, low molecular weight hJHBPs are limited to only Lepidoptera and Diptera (review: Trowell, 1992), they have, nevertheless, been characterized extensively because relatively large quantities of larval hemolymph can be easily obtained from just a few insects. The low molecular weight hJHBPs are usually monomeric and range in molecular weight from 25 to 35 kDa. Although low molecular weight hJHBPs have been identified in a number of lepidopterans, only those in B. mori, G. mellonella, and M. sexta have been extensively studied. 3.7.8.2.1. Chemical and physical characteristics The lepidopteran hJHBPs are monomers composed of approximately 220–240 amino acid residues. In species where the primary sequence of hJHBP has been deduced, four to six cysteine residues have been reported. Manduca sexta hJHBP, which has six cysteine residues (Park and Goodman, 1993) and G. mellonella hJHBP, which has four cysteine residues (Kolodziejczyk et al., 2001), contain two disulfide bridges. The results of CNBr cleavage, together with extrapolated data from the study of G. mellonella hJHBP, suggest that the disulfide bridges in M. sexta hJHBP are probably formed from the Cys9 to Cys16 and Cys151 to Cys195 residues. That the cysteine residues at position 9 and 16 are important to JH binding was demonstrated with the H. virescens hJHBP. This hJHBP displays a nearly identical alignment of cysteine residues with those in the M. sexta hJHBP (Wojtasek and Prestwich, 1995). Using the H. virescens hJHBP construct, Wojtasek and Prestwich (1995) generated mutant hJHBPs in which an alanine was substituted for a cysteine residue at each of the cysteine positions. They discovered that Cys9 and Cys16 are critical for JH binding but that cysteine residues at other sites are much less important. Park and Goodman (1993) demonstrated that of the two remaining cysteine residues not involved in disulfide bond formation, one is inaccessible to alkylating agents, implying an internal location within the protein. The other cysteine residue is easily modified, suggesting a location on the surface of the protein. The role of disulfide bridges in overall structure and hormone binding has yet to be determined. Initial studies indicated that the M. sexta hJHBP is not glycosylated (Goodman et al., 1978a); however, more recent evidence suggests hJHBPs from both

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G. mellonella and M. sexta are indeed glycosylated. Using matrix assisted laser desorption ionization time of flight (MALDI-TOF) measurements to determine the molecular mass of the native protein and to calculate the deduced amino acid residue mass, it has been determined that carbohydrates account for approximately 10% of the molecular mass of the M. sexta hJHBP (Goodman, unpublished data). Computer analysis of potential glycosylation sites indicates only one potential motif, that surrounding the Asn66 residue of the mature, secreted protein. The carbohydrate moiety can be enzymatically cleaved from the hJHBP using N-glycosidase F; however, hJHBP is resistant to cleavage by other glycosidases such as neuraminidase and endoglycosidase H (Goodman, unpublished data). Computer analysis of potential glycosylation motifs in the hJHBP from G. mellonella indicate multiple sites for glycosidation, including an O-GalNAc site (Thr116), N-glycosylation sites (Asn3, Asn93) and O-b N-acetylglucosamine (O-bGLcNAc) sites (Ser12, Thr116, Thr186) (Duk et al., 1996). Duk et al. (1996) confirmed this observation using lectin binding assays, gel staining, and mass spectrometry and suggested there are five major types of oligosaccharide chains, each containing two GlcNAc residues and two, three, or five mannose residues. The function of these carbohydrate moieties is unknown; however, glycosylation of hormone transport proteins in vertebrates is thought to limit proteolytic cleavage of the protein and thereby increase its half-life in the circulatory system (Westphal, 1986). While other potential posttranslational modifications can be predicted from computational programs, no experimental evidence for the modifications has been reported. The lepidopteran hJHBPs interact with their ligands at concentrations in the low nanomolar range (Goodman, 1990; Trowell, 1992; deKort and Granger, 1996); however, the homologs are not all bound with the same affinity. It was originally thought that the least polar of the homologs was bound with the highest affinity (Goodman et al., 1978b), following the polarity rule which states that within a series of small nonpolar homologous hormones, the least polar of the group will be bound with the highest affinity (Westphal, 1986). Park et al. (1993), in an extensive study of M. sexta hJHBP binding kinetics, demonstrated that the polarity of the JH homologs was not as important to the binding equilibrium as first proposed. This study revealed that the dissociation constants for the binding protein hormone complex are approximately the same for JH I or II, but as previously observed, the interaction with JH III is considerably weaker.

Although the polarity rule may not apply to the equilibrium constants, it is quite clear that the more polar the homolog, the shorter the half-life of the hormone–protein complex (Park et al., 1993). Thus, JH I, the least polar of the homologs tested, has the longest half-life (29 s), while JH III, the most polar homolog, has the shortest (13 s). These halftimes of dissociation are consistent with vertebrate hormone transport proteins (Mendel, 1989). The hJHBP of M. sexta preferentially binds the naturally occurring enantiomers of (10R)-JH III (Schooley et al., 1978b), (10R,11S)-JH I, and (10R, 11S)-JH II (Park et al., 1993). This enantiomeric specificity appears universal among the high affinity JH binding proteins, whether they are lipophorins or hexamerins (Trowell, 1992; deKort and Granger, 1996), indicating the binding site for the hormone is reasonably selective. In addition to binding studies using the JH homologs and their respective enantiomers, studies have been conducted using geometrical isomers of the hormones, hormone metabolites, and biologically active analogs (Goodman et al., 1976; Peterson et al., 1977, 1982). The results indicate that highly active JH analogs and JH metabolites do not interact with hJHBP. Armed with this information, several investigators have speculated on the nature of the ligand binding domain and its constituent residues. Goodman et al. (1978b) suggested the binding site may be a hydrophobic cleft on the surface of the protein. The primary interactions between the hormone and the binding site would occur along the alkyl side chains of the hormone and the methyl ester moiety at C1, which together form a distinct hydrophobic surface. Perturbation of that surface, especially by the introduction of highly polar groups at C1, significantly reduces binding. However, the site is not sterically hindered since nonpolar additions such as ethyl and propyl groups at the C1 position reduce binding only moderately (Peterson et al., 1977). Hydrogen bonding, which involves electrophilic residues in the binding site, potentially anchors the hormone at the epoxide and C1 positions and may be more important for interaction with JH III than JH I. Support for this idea comes from the studies of Prestwich and Wawrzen´ czyk (1985), who found that the interaction between the naturally occurring JH I enantiomer and hJHBP was only several fold greater than its antipode, suggesting that the binding site could not only accommodate the 10R,11S enantiomer but the 10S,11R enantiomer as well. In contrast, the significant binding energy derived from the (10R)-JH III epoxide–hJHBP interaction is probably disrupted in the (10S) configuration. Thus, the reduction in

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the hydrophobic ‘‘face’’ of JH III places considerably more importance on the hydrogen bonding between the epoxide and methyl ester of JH III and the hJHBP. Using photoaffinity labeling, Touhara and Prestwich (1992) attempted to map the binding site of the M. sexta hJHBP. On the basis of labeling patterns, they proposed that the region from Ala184 to Asn226, a region predicted to be a hydrophobic domain, is involved in the recognition of the lipophilic backbone of JH. Another region, Asp1 to Glu34, also appears to be involved, leading these investigators to propose a model in which the two regions are connected by disulfide bridges to form a two-sided hormone binding pocket. More recently, Tesch and Goodman (unpublished data) discovered that trypsin treatment of hormonesaturated (holo-hJHBP) and hormone-free hJHBP (apo-hJHBP) from M. sexta yields strikingly different results, raising questions about the Touhara and Prestwich model. When holo-hJHBP is treated with trypsin, the C-terminus of hJHBP from Lys180 onward is rapidly removed. The N-terminus (first 180 residues) of the molecule remains intact for approximately 1 h, despite the fact that it contains at least 10 well-documented tryptic cleavage sites. The trypsin-truncated hJHBP is still capable of binding JH, indicating that the C-terminus (last 46 residues) of the protein, despite its elevated level of hydrophobicity, is not directly involved in binding. Conversely, apo-hJHBP, when treated with trypsin, is completely destroyed within a few minutes. Similar findings were observed in G. mellonella (Wieczorek and Kochman, 1991). These results suggest that the binding site is not located in the C-terminus as proposed by the Touhara and Prestwich model but rather lies upstream, closer to the N-terminus. Rodriguez-Parkitna et al. (2002) hypothesize that the C-terminus may serve as a ‘‘barrel cover’’ that changes its position upon hormone binding. Thus, while the C-terminus may not be directly involved in hormone binding, it may have other important roles, such as target cell docking functions or hormone transfer to the cell. Hormone-induced conformational changes have been observed in certain of the vertebrate serum steroid hormone binding proteins (Grishkovskaya et al., 2002), and there is good evidence indicating that hJHBP undergoes a conformational change upon interaction with JH, but the degree of change appears to vary with the species. While the hormone appears to induce conformational changes that mask certain regions, no differences were detected between apo- and holo-hJHBP in sedimentation rate or electrophoretic migration studies in M. sexta

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(Tesch and Goodman, unpublished data). The studies of Touhara et al. (1993) on recombinant M. sexta hJHBP did not reveal differences in circular dichroism spectra of apo- and holo-hJHBP, indicating that the hormone does not induce changes in secondary structure. In contrast to these studies, Wieczorek and Kochman (1991) demonstrated a shift in the sedimentation coefficient between the apo-hJHBP (2.30S) and holo-hJHBP (2.71S) forms of the hJHBP from G. mellonella, as well as slight changes in electrophoretic mobility between these two forms. More recently, the Kochman group used circular dichroism analyses to confirm their earlier discovery that hormone binding leads to changes in the secondary structure of hJHBP in G. mellonella (Krzyzanowska et al., 1998). It is unclear why these changes in secondary structure were not detected in the M. sexta hJHBP. It may well be that small but significant differences between the primary and secondary structures of the M. sexta and G. mellonella hJHBPs are responsible. These initial studies on ligand-induced conformational change are highly intriguing but unfortunately offer only a glimpse into the structure and dynamic conformational changes. Crystallographic studies, such as those currently under way with the G. mellonella hJHBP (Kolodziejczyk et al., 2003) will provide the desired information on the nature of the JH binding site and ligand-induced conformational change in the protein. 3.7.8.2.2. Protein structures The high affinity, low molecular weigh hJHBPs that have been sequenced to date display a reasonable degree of identity in alignment (25%), but as noted by RodriguezParkitna et al. (2002), no one region shows a higher than average homology, thus precluding the identification of a putative JH-binding domain (Figure 14). These investigators examined other JH-associating proteins for potential sequence similarities and concluded that there was little similarity among the hJHBPs and ligand binding domains of potential nuclear receptors, insect transferrins, or the high affinity, high molecular weight JHBPs. From this analysis, it may be inferred that the composition and sequence(s) of amino acids residues making up the hydrophobic JH binding domain are not under extreme selective pressure, as long as the site is lined with hydrophobic residues that permit interaction with the aliphatic side chains of the hormone. However, the site must contain the hydrophilic residues to permit interaction with the epoxide moiety. As more protein sequences enter the data bases, a picture is emerging that suggests the lepidopteran

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Figure 14 Multiple alignment of the sequenced hemolymph JH binding proteins (hJHBPs) from Lepidoptera. Amino acid sequences of M. sexta (Madison wild-type) hJHBP (GenBank Accession no. AAF45309), B. mori hJHBP 2 (AAF19268), H. virescens hJHBP (AAA68242), and G. mellonella hJHBP (AAN06604). Alignment was performed using the CLUSTAL W algorithm. Letters in bold represent residues in the signal peptide. Symbols beneath the columns of amino acid residues represent the degree of conservation in each column: (*) indicates residues completely conserved; (:) indicates residues with only very conservative substitutions; (.) indicates residues with mostly conservative substitutions.

hJHBPs belong to a superfamily of proteins, the calycins (Rodriguez-Parkitna et al., 2002), which include fatty acid binding proteins, avidins, lipocalins (see Chapter 4.8), and metalloprotease inhibitors (Flower, 1996, 2000). Proteins in this family are typically low molecular weight, secreted proteins that bind small, hydrophobic molecules. Members of the calycin protein family share a b-barrel structure composed of 8 or 10 b-sheets that are flanked by a-helices in the N- and C-termini. While sequence similarity between the hJHBPs and other members of the superfamily is low, modeling of the G. mellonella and M. sexta hJHBP secondary structure indicates considerable similarity to insecticyanin and human plasma retinol binding protein, both well-studied lipocalins (Rodriguez-Parkitna et al., 2002). Given the structural information gleaned to date, it will not be surprising if in the near future, the lepidopteran hJHBPs are firmly placed in the lipocalin subfamily of calycins.

3.7.8.2.3. Genomic structures At the present time, the genomic structure of only one hJHBP, that from M. sexta (Madison wild-type strain), has been examined (Orth et al., 2003a). The hJHBP locus (GenBank Accession no. AF226857) consists of five exons spanning 6.7 kb and is flanked by typical eukaryotic and insect-specific transcriptional control elements. The first exon encodes the 50 untranslated region, a signal peptide plus the first two amino acids of the open reading frame, while the other exons encode the mature, secreted protein and the 30 untranslated region. The proximal promoter region contains a A/T-rich region similar in position, but slightly longer than that of the JH esterase gene from T. ni (Jones et al., 1998); however, both of these genes lack the canonical TATAA motif normally associated with the proximal promoter region. Computational analysis indicates numerous potential transcription factor binding sites in the 50 -flanking region of the gene. Most intriguing are

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two closely spaced ECR response elements (see Chapter 3.5) approximately 5 kb upstream from the start site. The JHs are presumed to act on the ecdysteroid signaling pathway (Riddiford, 1994), regulating gene expression by modulating the transcriptional activity of the ecdysteroid receptor and its heterodimeric partner, Ultraspiracle (Jones and Sharp, 1997) (see Section 3.7.12.2). Considering that both the protein concentration (Hidayat and Goodman, 1994) and the hJHBP gene expression (Orth et al., 1999) undergo a significant decline during a larval-to-larval molting period, it is reasonable to speculate that ecdysteroids, acting via these ecdysteroid response elements, may be involved in the regulation of hJHBP expression and may indirectly modulate JH titers. Recent studies reveal that strain-related polymorphisms in the hJHBP nucleotide sequence, outside the open reading frame, lead to significant differences in the titers of this protein. A comparison of hJHBP levels in two different strains of M. sexta revealed that one strain, the Seattle wild-type (Swt), has levels that are nearly 60% lower than those of the Madison wild-type strain (Mwt) (Young et al., 2003). The Swt strain exhibits an hJHBP allele, Snb, that differs significantly from the previously described Md allele found in the Mwt strain (Orth et al., 2003a) by the presence of a novel insertion in intron 2. The relatively small size of the insert (1 ng) inducing a downregulation of hJHBP mRNA abundance. Curiously, hormone application at other times during the stadium does not elicit this response, which has lead to the suggestion that JH-induced transcription of hJHBP is already maximally stimulated at these times and further elevation of hormone levels by topical application has a dampening response. Thus, the control of hJHBP levels appears to be tied to fluctuating JH titers during the intermolt period; elevated titers of the hormone downregulate hJHBP mRNA and protein, while reduced JH titers elevate both message and protein. During the molting period, the elevated levels of ecdysteroids required for molting reduce expression of the hJHBP gene. If the sole function of hJHBP is hormone delivery, then the developmental pattern and excess titer remain a mystery, but as with the vertebrate serum hormone binding proteins, the hJHBPs may be more than transport macromolecules. 3.7.8.3. Functions p0680

Facilitating the transport and dispersal of JH to distant target sites is the most obvious function of the hJHBPs, yet other equally important roles for the binding protein have been hypothesized or experimentally determined (Westphal, 1986; Goodman, 1990). These functions include: (1) reducing the levels of promiscuous binding to nontarget tissues; (2) reducing enzymatic degradation of the hormone (Hammock et al., 1975); (3) acting as a scavenger to enhance JH metabolism in larval hemolymph (Touhara et al., 1996) or the developing embryo (Orth et al., 2003b); (4) providing a peripheral reservoir of hormone that is readily available at the

target tissue; (5) enhancing JH synthesis in the CA by acting as a sink; and (6) aiding hormone movement from the hemolymph into the target cell. The first four functions are either well established or have at least gained a level of scientific credibility. Preliminary evidence now suggests that hJHBP may promote the synthesis of JH by dispersing the hormone from the immediate vicinity of the CA. Using the RCA to assess the effect of several JH-interacting proteins (bovine serum albumin, JHdirected antiserum, and M. sexta hJHBP) on JH biosynthesis in vitro, it was observed that only hJHBP significantly increased the incorporation of radiolabeled tracer into JH (Granger and Goodman, unpublished data). Manduca sexta CA synthesized 60% more JH when incubated in medium containing physiological levels of hJHBP. While the results are preliminary, they do suggest that the binding proteins aid the transit of JH from the CA and thus could prevent hormone from accumulating nonspecifically in the vicinity of the glands. The binding proteins, in essence, create a longer feedback loop and thus block short-circuiting of JH biosynthesis. The most intriguing unexplored aspect of the hJHBP functions is that of its action at the surface of a target cell. A considerable body of evidence has convinced most vertebrate endocrinologists that hormone binding proteins release their ligands into the circulatory system, thus allowing them to enter the cell in the unbound state (Westphal, 1986; Mendel, 1989; Grasberger et al., 2002). For some hormones, this is a reasonable conclusion; however, several protein-bound hormones and vitamins have specific cell surface receptors for their respective transport proteins (review: Kahn et al., 2002). Retinol binding protein (RBP), for example, binds to a plasma membrane receptor that participates in transfer of the ligand from the serum transport protein to the cytoplasmic retinol binding protein (Sundaram et al., 1998, 2002). As levels of apoRBP far exceed those of the holo-RBP, a conformational change in the protein must occur so that cell surface receptors are not interacting with ligand-free RBP (Chau et al., 1999). Even more striking are the roles of sex hormone binding globulin (SHBG), a vertebrate serum protein that binds and transports the sex steroids. As is the case for RBP, target tissues requiring the sex steroids display a plasma membrane receptor for SHBG. Unlike the RBP receptor, the SHBG receptor binds only the unloaded form of the transport protein; the hormone-loaded SHBG complex does not bind (Kahn et al., 2002). Sex steroids then bind to the receptor-bound SHBG on the cell surface which, in turn, activates adenylyl cyclase. The role of the

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The Juvenile Hormones

activated second messenger cascade remains unknown. The extracellular transport protein is more than just a vehicle for hormone dispersal; indeed, it may need to be anchored at the cell surface to facilitate uptake. Moreover, the receptor–protein complex may have its own intrinsic signaling ability. Whether this complex cascade of hormone transfer, cell surface receptors, and activation of second messengers occurs in insects remains to be investigated.

3.7.9. Catabolism of the Juvenile Hormones

p0705

The maintenance of effective hormone titers at the target site is a delicate balancing act among various processes: synthesis, delivery, metabolism, and cellular uptake. Endocrine signals, by their very nature, must be transitory to effect their exquisite control of the target response. Soon after the chemical structures of JHs were elucidated in the late 1960s, a search for hormone-inactivating enzymes began that has continued unabated to the present, with better than 350 articles describing some aspect of JH metabolism. This prodigious number of publications reflects the agricultural interest in targeting catabolism as a means of insect pest control, since it is assumed that dramatically increasing or decreasing catabolism of JH may lead to developmental derailment (Bonning and Hammock, 1996). While this novel means of pest control has yet to be commercially exploited, the biological information gleaned from studies on JH metabolism is of considerable importance in understanding hormone titer regulation. For earlier reviews of JH catabolism, the reader is referred to Hammock (1985), Roe and Venkatesh (1990), deKort and Granger (1996), and Gilbert et al. (2000). The initial step in JH catabolism may occur in the hemolymph or in target cells, and results in different products; however, the end point is the same: biological inactivation of the hormone. Hemolymph inactivation of JH is the result of enzymatic activity of several different esterases capable of hydrolyzing the hormone at the C1 position to form the metabolite, JH acid (Hammock, 1985; Roe and Venkatesh, 1990) (Figure 2). Although cellular inactivation of JH is primarily the result of epoxide hydrolases that hydrate the epoxide at the C10, C11 position to form the JH diol (Figure 2), cells also possess esterase activity that yields JH acid (Lassiter et al., 1995; Roe and Venkatesh, 1990). Our examination of JH catabolism begins with the hemolymph where esterases represent the primary mechanism of hormone inactivation.

357

3.7.9.1. Juvenile Hormone Esterases

The hemolymph of a number of species contains esterases that are capable of hydrolyzing the ester at the C1 position to form JH acid. While the early literature suggested that there are two classes of esterases responsible for JH hydrolysis, JH-specific esterase (JHE) and nonspecific or general esterases (Sanburg et al., 1975), recent work has been less clear about the role of the nonspecific enzyme(s) (Gilbert et al., 2000). General esterases were originally defined as those enzymes that could metabolize both a general substrate, a-naphthyl acetate, and JH (Sanburg et al., 1975) and were inhibited by the inhibitor O,O-diisopropyl phosphorofluoridate (DFP). More recent evidence suggests that the general esterases are less important in JH metabolism than first thought (Roe and Venkatesh, 1990; Gilbert et al., 2000). Because conversion to JH acid can be carried out by a number of enzymes, Hammock (1985) proposed a working definition for a JHE. From a biochemical standpoint, such an enzyme should have a low apparent Km for JH and it should therefore hydrolyze JH with a high kcat:Km ratio (see Section 3.7.9.1.2). Furthermore, the enzyme must be able to hydrolyze JH in the presence or absence of a JH binding protein. From a biological standpoint, the JHE must have activity that correlates with a decline in JH titer. The premise that these enzymes are vital to hormone titer regulation has thus received considerable attention. 3.7.9.1.1. Physical properties JH-specific esterases (EC 3.1.1.1) are members of the carboxylesterase family and have been studied in at least six orders of insects, including Thysanura, Orthoptera, Hemiptera, Coleoptera, Diptera, and Hymenoptera (Hammock, 1985); as molecular cloning becomes more widely used, more insect orders will undoubtedly be added to this list. To date, the best characterized of the JHEs are those from the hemolymph of Lepidoptera. The JHEs of Lepidoptera appear to be composed of a single polypeptide chain with a relative molecular mass of approximately 63–67 kDa (Roe and Venkatesh, 1990; Hinton and Hammock, 2003). The hemolymph of the coleopteran L. decemlineata, contains a JHE with a native Mr of 120 kDa; it appears to be composed of two polypeptide chains of approximately 57 kDa each (Vermunt et al., 1997a). A similar situation exists in the orthopteran Gryllus assimilis, where the native JHEs displays a Mr of 98 kDa and is composed of two identical subunits of 52 kDa (Zera et al., 2002). The

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358 The Juvenile Hormones

isoelectric points of all the JHEs studied are acidic, falling into the range of 5.2 to 6.1 (Abdel-Aal and Hammock, 1986; Hinton and Hammock, 2003). Isoelectric focusing of apparently pure JHE from several species yields two to four distinct proteins, suggesting heterogeneity in posttranslational modifications such as glycosylation. The JHE of the lepidopteran T. ni is glycosylated (Hanzlik and Hammock, 1987); however, the JHE of M. sexta lacks glycosylation, even though computational analysis of its sequence indicates three potential glycosylation sites (Kamita et al., 2003). It should be noted that even a baculovirus expression system containing a single JHE construct still yielded several isoforms that could not be explained by limited proteolysis or differential deglycosylation (Hinton and Hammock, 2003).

p0735

3.7.9.1.2. Kinetic parameters and specificity JHE kinetics and substrate specificity have long been a central focus of JHE studies, owing to the obvious agricultural potential of this information. Elucidation of JHE kinetic constants also permits predictions of the rate limits of JH metabolism, as well as provide valuable insight into to how peripheral JH levels are regulated. A number of reports from the 1980s indicate that among lepidopterans, the apparent Michaelis constant (Km) for the naturally occurring JHs ranged from 108 to 106 M (Roe and Venkatesh, 1990). JH titers in the species tested are at least 10–100 times lower than the estimated Km concentration, indicating that the enzyme is very sensitive to changes in JH concentration, but suggesting that its catalytic potential is wasted. Since the Km values seem unduly high with regard to substrate concentration, investigators have examined the individual rate components of the enzymatic reaction to better explain the observed data (Sparks and Rose, 1983; Hammock, 1985; Abdel-Aal and Hammock, 1986). In the simplest of terms, the velocity of JHE or any enzymatic reaction can be viewed as the sum of various rate components and the concentration of the reactants, and can thus be defined by the familiar equation: k1

k2

E þ S Ð ES * E þ P k1

½1

where E represents enzyme, S represents substrate, ES represent the Michaelis complex, and P the product. Under physiological conditions, where the concentration of JH is several orders of magnitude lower than the Km, the rate at which JH acid is formed is a function of JH interaction with JHE (k1 and k1) and the rate of product formation

(k2). Several important lessons can be drawn from the kinetic data for JHE. First, JHE has a very high affinity for JH. Second, it has a relatively low turnover number or kcat, where kcat is the maximum number of substrate molecules (JH) converted to product (JH acid) per active site per unit time. These factors make the enzyme a very effective scavenger that can ‘‘find’’ JH at physiological concentrations and convert it to JH acid (Abdel-Aal and Hammock, 1986). Substrate specificity of JHE represents another characteristic that seems counterintuitive. The Km and Vmax that the JHEs display towards the JH homologs is surprisingly low when compared to other substrates, such as a-naphthyl acetate. For example, recombinant M. sexta JHE displays a Km (410 mM) and Vmax (21 mmol min1 mg protein1) for a-naphthyl acetate that is considerably higher than for its natural substrate, JH III (Km ¼ 0.052 mM, Vmax ¼ 1.4 mmol min1 mg protein1) (Hinton and Hammock, 2003). As noted by Fersht (1985), when specificity is used to discriminate between two competing compounds, it should be determined by the kcat /Km ratios, and not Km alone. The kcat/Km ratio for JH III and a-naphthyl acetate are 27 and 0.04, respectively, underscoring the high degree of specificity of JHE for JH III (Hinton and Hammock, 2003). While these kinetic parameters suggest that the enzyme has a high degree of specificity, it appears that JHEs from several species also hydrolyze methyl and ethyl esters of JH I and III at very similar rates (Grieneisen et al., 1997). Moreover, even n-propyl and n-butyl esters of these homologs can serve effectively as substrates, albeit at a lower rate of hydrolysis. As might be expected, the unnatural (2Z,6E)-JH I ethyl ester isomer is not metabolized by JHEs from M. sexta or H. virescens. Since hJHBPs have a significant effect on substrate availability (Peter et al., 1979; Peter, 1990; King and Tobe, 1993), studies on JHE specificity must be performed with relatively pure preparations of the enzyme. In those few studies using highly purified or recombinant T. ni JHE, it was determined that the JHE hydrolyzed JH I more rapidly than JH II or III and that it displayed a faster catalytic rate for the naturally occurring enantiomer, (10R,11S)-JH II, than for the (10S,11R)-JH II form (Hanzlik and Hammock, 1987). However, a comparison of the kcat/Km ratios for the enantiomers indicated that hydrolysis of the two forms was equivalent (Hanzlik and Hammock, 1987). The same holds true for the gypsy moth, Lymantria dispar, in which the JHE shows no enantiomeric selectivity and appears unable to discriminate between

The Juvenile Hormones

p0750

homologs (Valaitis, 1991). One of the most unexpected groups of JHE substrates is found in the naphthyl and p-nitrophenyl series. While it was originally concluded that hemolymph JHE from some species, such as M. sexta, could not use a-naphthyl acetate as a substrate (Coudron, 1981), the JHEs of other species can recognize members of the naphthyl series (Rudnicka and Kochman, 1984; Hanzlik and Hammock, 1987). More recently it was demonstrated that recombinant JHE from M. sexta, H. virescens, and T. molitor can hydrolyze naphthyl compounds with chain derivatives eight carbons long, thus dispelling the long-held belief that JHE from M. sexta is unable to use naphthyl derivatives as substrates (Kamita et al., 2003). Another counterintuitive observation is that while the major role of JHE is the conversion of JH to JH acid, both native and recombinant JHEs from several species can, under the appropriate conditions, transesterify JH to form the higher ester homologs, i.e., JH ethyl, JH n-propyl, and JH n-butyl esters (Grieneisen et al., 1997). Although JHE-mediated JH transesterification may be a curiosity limited to the test tube, Debernard et al. (1995) demonstrated that when JH III, dissolved in ethanol (10 ml), was injected into L. migratoria, it was both converted to JH III acid and transesterified to the JH III ethyl ester. With regard to this particular study, it should be noted that care must be taken to avoid artifacts when alcohols are used as carrier solvents for the hormone in JHE assays. Nevertheless, while JHE in a biological milieu clearly serves as an esterase, these new findings imply that the enzyme may have other physiological roles (see Anspaugh et al. (1995) for other possible roles for JHE). 3.7.9.1.3. Protein structure and catalytic site While the catalytic action of JHE is probably similar to that of other esterases, there is some uncertainty about the molecular mechanism by which it operates and some question as to how it can effectively hydrolyze a chemically stable conjugated ester (Hinton and Hammock, 2003). A putative three-dimensional structure of the JHE of H. zea has been generated using homology modeling, a structure-building process that uses computer algorithms, based on the assumption that proteins with homologous sequences have similar three-dimensional structures (Thomas et al., 1999). Using the structures of two well-defined carboxylesterases (sequence identity 28%) as models, these investigators demonstrated that JHE belongs to the a/b hydrolase fold family. The H. zea JHE (GenBank Accession no AF037196) possesses a putative catalytic triad of amino acids, including the nucleophilic Ser224,

359

as part of the characteristic motif, G-X-S-X-G (X represents any amino acid residue). The second and third components to the triad are a base, His465, which is part of the G-X-X-H-X-X-D/E motif , and an acid, Glu353, that lie in a deep cleft of approximately 23 A˚ . The importance of these residues to JH catalysis was confirmed through site-directed mutagenesis (Ward et al., 1992). In addition to the catalytic site, the cleft is lined with a number of hydrophobic amino acids, a situation that might be expected, considering the hydrophobic nature of JH. One might reasonably expect the catalytic site of JHEs of other species to show some sequence similarity and, not surprisingly, they do. A comparison of sequences among the known lepidopteran JHEs, those of B. mori (Hirai et al., 2002), Choristoneura fumiferana (Feng et al., 1999), H. virescens (Hanzlik et al., 1989), and M. sexta, (Hinton and Hammock, 2001), reveals better than 30% identity. The residues associated with the catalytic site are in complete agreement and display the appropriate alignment. Even in species outside the order, such as D. melanogaster (Campbell et al., 1992) and T. molitor (Hinton and Hammock, 2003), the catalytic sites align with those in Lepidoptera (Feng et al., 1999; Kamita et al., 2003). The only species that does not show alignment of the putative catalytic site is L. decemlineata (Vermunt et al., 1997b), which is surprising given the consensus displayed by another beetle, T. molitor. With the exception the putative catalytic site residues and a sequence surrounding Ser224, the JHEs as a family are not highly conserved. This might be expected, however, since only the cleft into which the substrate fits needs to be lined with hydrophobic residues (Kamita et al., 2003). 3.7.9.1.4. Juvenile hormone esterase inhibitors The development of effective JHE inhibitors has been an ongoing process since the first studies more than 25 years ago. While JHE inhibitors have yet not been commercialized for agricultural use, they have, nevertheless, been important tools in the discovery of the role of JH catabolism in vivo. Two important series of JHE inhibitors have emerged from these studies, the trifluoromethyl ketones (TFKs) and the phosphoramidothiolates (Roe and Venkatesh, 1990). The TFKs, such as 1,1,1-trifluorotetradecan-2-one (TFT), were initially suspected to act as transitional state analogs, mimicking the a,b saturation of JH (Hammock et al., 1982) (Figure 15). The transitional state, where the enzyme and substrate form a complex, represents a transitory period in which the chemical bonds of the substrate

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360 The Juvenile Hormones

since they are highly specific and appear to have little effect on nonspecific esterases (Roe et al., 1997). In contrast to the TFKs, the organophosphate inhibitors of JHE, most importantly O-ethyl S-phenyl phosphoramidothiolate (EPPAT) (Figure 15), bind irreversibly to JHE (Hammock, 1985). Like the TFKs, EPPAT appears to be highly selective for JHE and can be used in vivo. While EPPAT is only a moderately effective inhibitor of JHE, it has a much longer life in vitro than the TFKs.

Figure 15 Chemical structures of JH III, JH esterase and JH epoxide hydrolase inhibitors. TFT, 1,1,1-trifluorotetradecan-2one; OTFP, 3-octylthio-1,1,1-trifluoro-2-propanone; OTPdOHsulfone, 1-octyl[1-(3,3,3-trifluoropropan-2,2-hydroxy)] sulfone; EPPAT, O-ethyl S-phenyl phosphoramidothiolate; MEMD, methyl 10,11-epoxy-11-methyldodecanoate; NOPU, N-[(Z )-9-octadecenyl]-N 0 -propyl urea.

are in the process of being broken and made. Thus, a transitional state analog has the properties of the unstable intermediate. In the catalytic site, the electron-withdrawing inhibitor increases the electrophilicity of the carbonyl group, enhancing its susceptibility to nucleophilic attack by the reactive serine (Hammock et al., 1982). While it was thought that mimicking the JH backbone was important to recognition, further modifications of TFT to resemble JH structure did not improve JHE inhibition. However, introducing a thioether beta to the carbonyl 3-octylthio-1,1,1-trifluoro-2-propanone (OTFP) (Figure 15) increased the potency of inhibition at least 100-fold (Abdel-Aal and Hammock, 1985). More recently, it has been demonstrated that 1-octyl[1-(3,3,3-trifluoropropan-2,2-dihydroxy)] sulfone (OTPdOH-sulfone) is a better inhibitor of JHE than OTFP (Roe et al., 1997). The increased inhibition displayed by OTFPsulfone stems from the increased hydration (Roe et al., 1997; Wheelock et al., 2001). The attractiveness of the TFK-containing inhibitors lies in their reversible binding of the enzyme, which permits their use in affinity-matrix purification of JHEs. Moreover, the TFKs are useful for in vivo studies

3.7.9.1.5. Genomic structures At the present time, the genomic structures of only two JHEs are completely known, those of H. virescens (GenBank Accession no. J04955) (Hanzlik et al., 1989; Harshman et al., 1994) and D. melanogaster (GenBank Accession no. AF304352) (Campbell et al., 2001). The JHE locus of H. virescens is 8 kb in length and is composed of five exons that give rise to an open reading frame of 1.8 kb (Harshman et al., 1994). Unfortunately, nothing is known about the 50 upstream region in this gene, although the core promoter region for JHE has been elucidated in another noctuid, T. ni (Jones et al., 1998). In D. melanogaster, the JHE is localized on the second chromosome and covers approximately 2.5 kb in region 52F12 (FlyBase). The gene is composed of five exons (note: Flybase incorrectly indicates six exons) that contain an open reading frame of 1.7 kb. In the 50 position immediately adjacent to the JHE gene is another carboxylesterase gene (Flybase CG 8424) that displays 42% identity with JHE and contains the putative catalytic sites (Kamita et al., 2003). This gene is nearly identical in length and is suggested to be a gene duplication (Campbell et al., 2001). Just downstream from JHE is the gene spin, which is reported to be implicated in female behavior (FlyBase CG8428). Since gene expression may be coordinately regulated by domain control elements (Spitz et al., 2003), a knowledge of neighboring genes may provide new insights into the regulation of JHE and potential molecular cascades in which the gene is involved. 3.7.9.1.6. Regulation The importance of JHE in regulating JH titers has prompted a number of investigators to examine its physiological and genetic regulation (review: Gilbert et al., 2000). The primary site of synthesis for the hemolymph JHEs, the fat body (Whitmore et al., 1974; Hammock et al., 1975; Wing et al., 1981), has been the primary focus of many studies that have been reviewed previously in some detail (Roe and Venkatesh, 1990; Roe et al., 1993; deKort and Granger, 1996; Gilbert et al., 2000). Tissues other than the fat body can also

p0775

The Juvenile Hormones

express the enzyme (see Roe and Venkatesh, 1990 for a review of the different tissues; Feng et al., 1999). Even the CA have been found to have JHE activity in a number of species (Sparagana et al., 1984; Wisniewski et al., 1986; Meyer and Lanzrein, 1989a, 1989b), and in one case, the glands secrete measurable, and developmentally fluctuating, amounts of the enzyme into incubation medium (Sparks et al., 1989; Janzen et al., 1991). Most studies conclude that the JHEs of different tissues and at different developmental time points are similar, if not identical, to the hemolymph form of JHE; however, their regulation may be different (Jesudason et al., 1992). Sparks et al. (1989) made the novel suggestion that the multiple isoforms of hemolymph JHE may be the result of its production and release into the hemolymph from a variety of tissues. To best understand JHE regulation, a brief description of hemolymph JHE levels is in order. In M. sexta, a peak in hemolymph JHE is observed prior to ecdysis in larval stadia two through five (Roe and Venkatesh, 1990), with activity in each stadium increasing in direct proportion to larval weight (Roe et al., 1993). Although levels of JHE activity during the earlier stadia are lower, the precise timing and appearance of JHE and its developmental profile in relation to total hemolymph protein suggests specific regulation. However, the role of the JHE in the early instars is unclear, since hemolymph JH titers remain relatively high even in the presence of enzyme (Hidayat and Goodman, 1994).

361

In contrast to the earlier stadia, the developmental profiles of JHE activity and of JH titers during the last stadium of M. sexta reveal two peaks (Vince and Gilbert, 1977; Baker et al., 1987; Roe and Venkatesh, 1990). Titers of JH are highest in the hours immediately following the molt from the fourth to the fifth stadium (Figure 16). They then drop dramatically, becoming undetectable by 48 h after ecdysis. JH titers rise a second time to peak at 156 h and then decline prior to metamorphosis. JHE activity begins to climb midway through the feeding period and peaks shortly before wandering. A smaller rise in JHE activity appears after the second peak of JH but drops at pupation (Baker et al., 1987). Similar developmental profiles are seen in the last stadium of S. littoralis (Zimowska et al., 1989) and L. decemlineata (deKort, 1990). Most studies have pinpointed the fat body as the primary source of JHE in the hemolymph (Wing et al., 1981; Wroblewski et al., 1990). However, Jesudason et al. (1992) suggest that the first peak of JHE, which occurs during the feeding stage, is of fat body origin, while the second peak, during the prepupal phase, is not. Alternatively, it could be that in this study, the preparation of prepupal fat body samples in the absence of protease inhibitors led to the destruction of JHE at this stage. While the JHE profile in hemolymph appears to be similar for most lepidopterans, it appears to differ outside of this order: some species have a single burst of JHE activity that declines either midway through the last stadium or at the very end of larval life (review: Roe and Venkatesh, 1990).

Figure 16 Hemolymph JH titer and JHE activity during the fifth stadium of M. sexta larval development. Closed circles, total titer for all homologs detected (JH 0, I, II, III). JH acids are not included. JH titers were combined for both females and males and expressed as the average. Closed squares, JHE activity. JHE activities were combined for both females and males and expressed as the average. W, onset of wandering behavior; P, time of pupal ecdysis. (Adapted from Baker, F.C., Tsai, L.W., Reuter, C.C., Schooley, D.A., 1987. In vivo fluctuation of JH, JH acid, and ecdysteroid titer, and JH esterase activity, during development of fifth stadium Manduca sexta. Insect Biochem. 17, 989–996.)

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362 The Juvenile Hormones

As can be seen in Figure 16, the rise in JHE activity after the JH titer drops does not support the concept that the enzyme is directly responsible for the decline in JH titers. Nevertheless, the enzyme displays exceptional scavenging properties, and it may be that the increase in enzyme activity occurs to ensure that no detectable JH is present during commitment to pupation. More problematic is the observation that JH is not totally eliminated when JHE is present. For example, a comparison of JH levels in the penultimate stadium of M. sexta (Hidayat and Goodman, 1994) with JHE levels during this same period (Roe and Venkatesh, 1990) reveals measurable JH in the presence of JHE. The results of similar studies of the fifth stadium in this species strongly suggest that the pre-wandering peak of JHE has no functional relationship to metamorphosis (Browder et al., 2001). While not stated explicitly, the same conclusion can be drawn from studies in T. ni (Sparks et al., 1979; Hanzlik and Hammock, 1988; Jones et al., 1990b), and in still another case, JH levels appear refractory to JHE in diapausing larvae of the European cornborer, Ostrinia nubilalis (Bean et al., 1982). Although hemolymph JHE levels follow a developmental pattern that suggests the enzyme may be directly involved in JH catabolism, there remains a degree of uncertainty about the assumed role of JHE in eliminating the hormone from the circulatory system. That JHE levels do not follow a developmental pattern typical of other hemolymph proteins prompted a search for possible regulatory mechanisms modulating its activity. A logical candidate for JHE regulation is JH itself, but regulation of JHE by JH in lepidopteran larvae, especially during the last larval stadium, depends on whether the insect is in the pre-wandering or post-wandering phase. In the pre-wandering phase of the last larval stadium, JH regulation of hemolymph JHE levels is ambiguous. The brain appears to play an inhibitory role in control of JHE activity, and there are other factors, such as tissue competence, that seem to be involved as well (Venkatesh and Roe, 1988; Jesudason et al., 1992; Roe et al., 1993). In contrast, manipulation of JH titers by surgical and chemical means during the post-wandering period of the last stadium has a direct impact on JHE activity (Roe et al., 1993; review: Gilbert et al., 2000). Treatment of larvae with JH or JH analogs leads to an increase in JHE activity in the hemolymph, while allatectomy leads to a decline in JHE that can be reversed by supplying exogenous hormone. At the molecular level, nuclear run-on experiments using the fat body of T. ni have confirmed that exogenous JH or JH analogs stimulate JHE expression and that the response can be

detected within 3 h of treatment (Venkataraman et al., 1994). Using a cell line derived from the midgut of C. fumiferana, Feng et al. (1999) demonstrated that JH can increase the abundance of JHE mRNA within 1 h, suggesting a potential upregulation of JHE expression by JH. However, these investigators rightfully caution that the apparent rise in message could also be due to stabilization of JHE mRNA. Curiously, the potent protein synthesis inhibitor, cyclohexamide, mimicked the action of JH, prompting the suggestion that an inhibition of protein synthesis leads to an increase in the accumulation of JHE message or, alternatively, stabilization of the message. In contrast to the apparent stimulation of expression levels by JH, the presence of 20E in the medium leads to a decrease in JHE mRNA, in a dose-dependent manner (Feng et al., 1999). This discovery indicates that regulation of JHE expression is more complex than first thought and that ecdysteroids must be considered in the overall regulatory system. The endocrine system is an important bridge between the target cell and the environment, and it is axiomatic that environmental factors are major players in regulation of the endocrine system. Factors such as stress (Gruntenko et al., 2000), circadian cues that induce diapause (Bean et al., 1982, 1983), nutrition (Cymborowski et al., 1982; Sparks et al., 1983; Venkatesh and Roe, 1988; Roe and Venkatesh, 1990), and parasitism (Hayakawa, 1990; see Edwards et al., 2001 and references therein) are all thought to play a role in regulating JHE levels and are undoubtedly transduced by the nervous system. 3.7.9.1.7. Catabolism of juvenile hormone esterase After a rapid increase midway through the feeding period of the last stadium in Lepidoptera, JHE levels drop dramatically immediately prior to the onset of wandering. Experimental evidence corroborates this observation: the half-life of recombinant JHE when injected into second instars of M. sexta is approximately 20 min, while other exogenously supplied proteins of similar molecular mass display half-lives measured in days (Ichinose et al., 1992). Since no cleavage products of JHE can be detected in the hemolymph, Ichinose et al. (1992) suggested that the enzyme is removed from the circulatory system by cellular uptake. It was discovered that the pericardial cells were responsible for uptake and destruction of the hemolymph JHE (Ichinose et al., 1992). Pericardial cells are a collection of cells that surround the insect heart in various configurations, depending upon the order (Wigglesworth, 1972; Crossley, 1985). In M. sexta,

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the pericardial mass is punctuated with numerous labyrinthine channels that increase exposure of the cells to the hemolymph (Brockhouse et al., 1999), and it is thought that this tissue mass is responsible for the removal of hemolymph proteins and small colloidal particles from the hemolymph (Crossley, 1985). It is speculated that the rapid removal of JHE from the hemolymph by the pericadial cells occurs via receptor-mediated endocytosis, followed by transport to lysosomes for destruction (Bonning et al., 1997). During the passage from the endosome to the lysosome, JHE interacts with putative heat shock cognate proteins with relative molecular masses of 100 and 140 kDa. These pericardial cellspecific proteins cross-react with a universal antiserum directed towards heat shock protein 70, hence the designation as cognate (Shanmugavelu et al., 2000). In addition to the heat shock proteins, a novel 29 kDa protein, termed P29, appears to bind and target JHE for lysosomal destruction. This protein has been detected in fat body and pericardial cells of insects from all five larval stadia in M. sexta. The P29 gene has been identified using a phage display library and has been sequenced (GenBank Accession no. AF233526). 3.7.9.2. Juvenile Hormone Epoxide Hydrolases

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The preponderance of literature on JH catabolism has focused on the role of JHE in metamorphosis, yet the less well-studied pathway, i.e., epoxide hydration via JH epoxide hydrolase (JHEH), may actually be more biologically relevant in some species (Gilbert et al., 2000). For example, when JH I is injected into early fifth instars of M. sexta, the major metabolite is not JH acid or JH acid-diol, but a JH diol phosphate conjugate (Halarnkar et al., 1993). Further evidence that JHEH plays a significant role in the reduction of JH titers at critical times can be found in the mosquito Culex quinquefasciatus, in which JHEH activity is two to four times higher than JHE activity throughout most of the life cycle (Lassiter et al., 1994). Moreover, two peaks of JHEH are seen during the last stadium, suggesting that the enzyme is involved in JH metabolism during critical developmental periods (Lassiter et al., 1995). Even in T. ni, with which so many of the JHE studies have been done, JH diol is a major metabolite or intermediate (Kallapur et al., 1996). It appears that the contribution of JHEH to the overall catabolism of JH varies extensively, even within the same order (Hammock, 1985). 3.7.9.2.1. Physical properties The JHEHs (EC.3.3.2.3) belong to a large family of proteins

363

that display the ab hydroxylase fold (Debernard et al., 1998) and they are responsible for hydration of the JH epoxide to a diol. Since hydrolases are involved in a wide range of enzymatic reactions and utilize multiple substrates, it has been justifiably cautioned that JHEHs may not be as specific as they are portrayed (Harris et al., 1999). Nevertheless, the term JHEH will be employed as it is commonly used in the literature. A number of species have been catalogued as having JHEH activity (Hammock, 1985), but in only a few species have the JHEHs been well characterized. Because a single species may have more than one form of JHEH (Harshman et al., 1991; Keiser et al., 2002), it has been suggested that the different JHEHs represent tissue-specific enzymes (Harshman et al., 1991). Epoxide hydrolases are present in both soluble (cytoplasmic) and insoluble (microsomal) forms, but the EHs responsible for hydration of the epoxide are found associated mainly with the microsomal fraction (Wisniewski et al., 1986; Touhara and Prestwich, 1993; Wojtasek and Prestwich, 1996; Harris et al., 1999; Keiser et al., 2002). JHEH appears to be a monomer, with a relative molecular mass ranging from 46 to 53 kDa (Harshman et al., 1991; Touhara and Prestwich, 1993; Harris et al., 1999; Keiser et al., 2002). In general, the pH range for JHEH activity is broad, extending from approximately pH 5 to 9, probably reflecting the involvement of the reactive histidine (pKa 6.5 for the imidazole group) in the catalytic site (Debernard et al., 1998). The JHEHs of D. melanogaster are heat- and organic solvent-tolerant, withstanding incubation at 55  C and concentrations of ethanol exceeding 40% (Harshman et al., 1991). Moreover, the recombinant enzyme from M. sexta retains full activity in the presence of low levels of the reducing agent dithiothreitol and a sulphydryl modifying reagent, iodoacetamide (Debernard et al., 1998). 3.7.9.2.2. Kinetic parameters, specificity, and the catalytic site Determinations of the kinetic parameters of the JHEH from the microsomal fraction of M. sexta eggs revealed Km values for JH I, II, and III of 0.61, 0.55, and 0.28 mM, respectively (Touhara and Prestwich, 1993). Interestingly, the Vmax:Km ratio for JH III, the least abundant egg homolog (Bergot et al., 1981a), was 25 times greater than that for JH I, suggesting that the enzyme displays considerably more specificity for JH III than the higher homologs (Touhara and Prestwich, 1993). This observation was confirmed by Debernard et al. (1998), using recombinant JHEH from M. sexta; while the enzyme could hydrolyze

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JH I, II, and several synthetic substrates, JH III was the most favored substrate. JH III is also a better substrate for JHEH than JH acid (Touhara and Prestwich, 1993; Kallapur et al., 1996), and in D. melanogaster, JH III is a more suitable substrate than JHB3 (Casas et al., 1991). If JH III is the optimal substrate for JHEH, then JHEH may indeed be the enzyme responsible for intracellular JH catabolism, especially since with the exception of JHB3, JH III is the only or predominant JH in all insect orders but Lepidoptera (see Section 3.7.2.1). However, as noted by Harris et al. (1999), more direct evidence, such as a correlation between inhibition of the enzyme in vivo and a decrease in JH metabolism, is required before assuming the enzyme is JH III-specific. As noted by several groups, the high degree of conservation of the catalytic site suggests that the mechanism of epoxide hydration is similar to that found in mammals (Roe et al., 1996; Wojtasek and Prestwich, 1996; Debernard et al., 1998). Using the JHEH of M. sexta as a model, Debernard et al. (1998) proposed a two-step catalytic process. The first step involves a nucleophilic attack of the epoxide at the least hindered position, C10, by Asp227. This, in turn, opens the ring, leading to the formation of an hydroxyl-alkyl enzyme. The neighboring Trp228 is thought to activate the epoxide for a nucleophilic attack (Harris et al., 1999). The second step involves the hydrolysis of this covalent intermediate by a water molecule that is activated by His428 and Asp350. The histidine residue activates water with an Asp or Glu residue acting as the proton scavenger from the histidine to reactivate the enzyme. The resulting product is (10S,11S)-JH diol. A variation on this putative reactive site was found by Linderman et al. (2000), in which the site uses Glu403 instead of Asp350 as the charge relay partner with the histidine residue. There is remarkable sequence conservation of the catalytic triad and its surrounding residues among both insects and mammals. All JHEHs, like their mammalian counterparts, contain a tryptophan residue at positions 150–155, forming part of the oxyanion hole that may stabilize the hydroxyl-alkyl intermediate (Lacourciere and Armstrong, 1993; Wojtasek and Prestwich, 1996; Keiser et al., 2002). 3.7.9.2.3. Juvenile hormone epoxide hydrolase inhibitors In contrast to the early and successful discoveries of highly effective JHE inhibitors, the search for JHEH inhibitors has been less productive until recently (Hammock, 1985; Casas et al., 1991; Harshman et al., 1991; Roe et al., 1996). Investigations have taken one of two directions, examining

either compounds that mimic the JH backbone (Roe et al., 1996; Linderman et al., 2000) or compounds based on urea and amide pharmacophores that are not subject to metabolism through epoxide degradation (Severson et al., 2002). Roe et al. (1996) demonstrated that methyl 10,11-epoxy-11-methyldodecanoate (MEMD), a long-chain aliphatic epoxide (Figure 15) displays an I50 (molar concentration needed to inhibit 50% of enzyme activity) in the low nanomolar range. A promising group of MEMD analogs was investigated by Linderman et al. (2000), who re-examined the structure–activity relationship of MEMD epoxide substitution and enantioselectivity. Two classes of MEMD analogs were synthesized, a glycidolester series and an epoxy-ester series. As a group, the glycidol-esters were more potent inhibitors than the corresponding epoxy-esters, by an order of magnitude. The inhibitory activity in both classes was found to be dependent upon the absolute configuration of the epoxide at C10, with the R configuration displaying the higher degree of inhibition. The inhibitory activity of the most potent compound of the series (I50 ¼ 1.2  108 M) is thought to be due to the hydroxyl group in the active site, which forms an additional hydrogen bond. This bond may stabilize the enzyme–inhibitor complex by inducing a conformational change, or it could reduce the rate at which the diol product dissociates from the enzyme’s active site. The other class of JHEH inhibitors is the analogs of the ureas and amide pharmacophores that have been demonstrated to be potent inhibitors of mammalian soluble and microsomal epoxide hydrolases in mammals (Severson et al., 2002). To date, none of the nearly 60 compounds tested are as active as the glycidol-esters in the inhibition of recombinant M. sexta JHEH (Figure 15), a surprising result, given their action on mammalian enzymes. The most potent of the series, N-[(Z)-9-octadecenyl]-N0 propyl urea (NOPU), has an I50 ¼ 8.0  108 M. Severson et al. (2002) suggest that when the inhibitor enters the catalytic site, the carbonyl group of the urea interacts with two tyrosine residues in the oxyanion hole. Since the inhibitor lacks an epoxide, it does not covalently bind to the reactive Asp227 residue, but it does block the catalytic site from further activity. 3.7.9.2.4. Genomic structures At the present time, the genomic structure of only one insect JHEH gene is known, that of D. melanogaster. The gene cluster, approximately 8.6 kb, is located on chromosome 2 in region 55F7-8, near the JHE locus at 52F1. As suggested by Harshman et al. (1991) and confirmed

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through genomic studies (FlyBase), there are multiple forms of JHEH in D. melanogaster. Annotation of the JHEH locus identifies three putative genes that display approximately 37% identity in their amino acid sequence. The genomic region containing JHEH1 (GenBank Accession no. NM_137541; CG15101) is approximately 1.8 kb and contains four exons. Approximately 0.7 kb downstream from JHEH1 lies JHEH2 (GenBank Accession no. NM_137542; CG15102), which is 2.2 kb in length and has two potential transcripts. One transcript has four exons (GenBank Accession no. NM_137524; CG15102) while the other has three (GenBank Accession no. NM_176233; CG15102). At 0.3 kb downstream from JHEH2 lies the gene Bari which encodes a transposable element. JHEH3 (GenBank Accession no. NM_137543; CG15106) lies about 0.3 kb downstream from the transposable element and is composed of three exons. No genes of obvious relation to JH action or catabolism lie close to the JHEH locus. JHEH1 and 2 are more related to each other than to JHEH3; all three putative genes display the residues of the catalytic triad in the correct sequence, and all are of about the same length. Computational analysis using CLUSTALW indicates that JHEH1 and 3 are most divergent in their first 70 residues, which might be expected if the N-terminal is needed for attachment to cell membranes. Since JHEH activity has been demonstrated in other cell fractions besides microsomes (Casas et al., 1991; Harshman et al., 1991), it may well be that the first exons of JHEH1 and 3 code for sequences that target these enzymes for different locations within the cell. Recent evidence suggests that the N-terminal of EHs may play another role. The soluble human EH, EPXH2, also possesses phosphatase activity, so the enzyme can not only transform epoxy fatty acids to their corresponding diols but also dephosphorylate dihydroxy lipid phosphates (Newman et al., 2003). The phosphatase activity localized to the N-terminal domain is unaffected by a number of classic phosphatase inhibitors. Alignment of EPXH2 with several of the insect JHEHs shows less than 25% amino acid sequence similarity under nonstringent conditions, and regions of similarity are limited to the C-terminal domain. Nevertheless, the possibility of an insect JHEH with phosphatase activity presents an interesting twist in the metabolic pathway for JH catabolism (see Section 3.7.9.3). 3.7.9.3. Secondary Metabolism of Juvenile Hormone: Juvenile Hormone Diol Kinase p0870

Secondary metabolism of JH has been examined in a number of investigations (Roe and Venkatesh, 1990;

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Halarnkar et al., 1993; Grieneisen et al., 1995), but as noted by Halarnkar et al. (1993), the results of these studies may be misleading, since the enzymes used may have contained multiple hydrolytic activities. In only one instance has an actual JH conjugate been unequivocally identified. The conjugate, (10S,11S)-JH diol phosphate (Figure 1), is the product of a two-step enzymatic process: conversion of JH to JH diol and then addition of a phosphate group to C10 (Halarnkar et al., 1993). The enzyme responsible for the phosphorylation of JH diol is JH diol kinase (JHDK), which has been characterized from the Malpighian tubules of early fifth instars of M. sexta (Grieneisen et al., 1995; Maxwell et al., 2002a, 2000b). The discovery of JHDK (EC 2.1.7.3) was made when an analysis of JH I metabolites in vivo yielded, in addition to JH diol and JH acid, a very polar JH I conjugate that was subsequently identified as JH I diol phosphate (Halarnkar and Schooley, 1990). 3.7.9.3.1. Physical properties JHDK from M. sexta Malpighian tubules is a cytosolic protein composed of two identical subunits of 20 kDa, as determined by mass spectrometry (Maxwell et al., 2002a). Gel filtration studies indicate it has a molecular mass of approximately 43 kDa, and it has been suggested that the native form of the enzyme is homodimeric. JHDK displays a Km in the nanomolar range for JH I diol, which is appropriate for an enzyme responsible for clearance of a hormone whose titers rarely exceeds 10 nM. Most significantly, the developmental profile of catalytic activity for JHDK parallels that for JHEH, a requisite if JH diol phosphate is a legitimate terminal metabolite. Analysis of the kcat/Km parameter for the diols of JH I, II, and III indicates that JH I diol is the preferred substrate, suggesting a preference for an ethyl group at the C7 position. JHDK requires both Mg2þ and ATP for activity (Grieneisen et al., 1995; Maxwell et al., 2002a), although excess Mg2þ or Ca2þ inhibits its activity (Maxwell et al., 2002a). When stored in the appropriate buffers, the enzyme is reasonably stable but it is very sensitive to various metal ions. The specificity of JHDK for JH I diol is relatively high, considering the multitude of potential phosphate acceptor groups present in a cell. The enzyme does not recognize methyl geranoate diol (one isoprenyl unit shorter than JH) nor methyl geranylgeranoate diol (one isoprenyl group longer than JH), yet it does recognize JH I ethyl ester diol. It also recognizes both JH diol enantiomers, indicating that absolute stereospecificity of the hydroxyl groups is of minor importance.

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Most surprising is the enzyme’s inability to recognize JH acid diols. If JH acid diol cannot be phosphorylated by JHDK, the fact that JH diol phosphate is a significant metabolite (Halarnkar et al., 1993) must be explained otherwise. JH acid diol could undergo further catabolism by cytochrome P-450s (Sutherland et al., 1998). However, the role of cellular JHE becomes problematic if the pathway catalyzed by JHEH and JHDK is the major pathway for JH catabolism in the cell. There may yet be undiscovered enzymes in cellular JH catabolism. 3.7.9.3.2. Genomic structures The sequence and hypothetical structure of the M. sexta and D. melanogaster JHDK genes have recently been examined by Maxwell et al. (2002b). A partial characterization of JHDK from whole body homogenates of D. melanogaster indicates that it is similar to the enzyme in M. sexta, with the exception of its subunit structure. The active D. melanogaster JHDK is composed of a single monomer of 20 kDa, while the active M. sexta JHDK is composed of two identical 20 kDa subunits. This latter gene (GenBank Accession no. AJ430670) has been sequenced and found to code for an enzyme that has 59% sequence identity and >80% similarity to sarcoplasmic calcium-binding protein 2 (dSCP2) of D. melanogaster (GenBank Accession no. AF093240; CG14904). Similarities in chromatographic properties, isoelectric point, and enzyme activity led Maxwell et al. (2002b) to conclude that dSCP2 is the probable D. melanogaster homolog of M. sexta JHDK. Maxwell et al. (2002b) used computer modeling and docking programs to generate a three-dimensional model of JHDK. They capitalized on two facts: (1) the catalytic site of JHDK must contain both a purine binding site (MgATP or MgGTP) and a hydrophobic pocket for JH diol; (2) the scaffolding for another sarcoplasmic calcium-binding protein is known. Both the M. sexta and D. melanogaster JHDKs contain the three conserved nucleotide-binding elements common to nucleotide binding proteins, surrounding the putative substrate-binding site. The model further demonstrates that the protein contains four domains that form two pairs of a helix–loop–helix motif (EF-hand) (Branden and Tooze, 1999). The model’s charge interactions in the hydrophobic binding pocket, as ˚ ), are complementary to the well as its depth (19 A extended conformation of the diol. Moreover, the hydrophobic nature of the binding pocket complements the C1 ester of the substrate and supports the

observation that JH diol is the only substrate for this enzyme (Maxwell et al., 2002a). 3.7.9.4. Juvenile Hormone Catabolism and New Directions

The field of JH catabolism is changing with the application of sophisticated analytical tools and the use of metabolism studies in vivo to uncover potential catabolic pathways. The combination of the two approaches has led to the surprising discovery that JH diol phosphate conjugates are major JH catabolites, thus emphasizing the role of JHEHs in clearing JH from the body (Halarnkar et al., 1993; Gilbert et al., 2000). That discovery has revealed a significant problem in the JHE phase-separation assay. While this assay is a powerful tool, it also detects polar metabolites resulting from enzymatic activity other than that of JHE. In addition, it is critical that the correct controls be employed. This point is driven home by the fact that JHEs from several species can, under the appropriate conditions, transesterify JH (Debernard et al., 1995; Grieneisen et al., 1997). Thus, the JHE phase separation assay should be employed only as the first step in the search for new JH catabolites. The assay should be followed by the use of the advanced chromatographic tools and detection systems now available for identifying trace polar metabolites. The need to demonstrate biological relevance applies not only to the newly discovered JH-like molecules (see Section 3.7.3.1), but also to the enzymes involved in JH catabolism. It may well be that some of the enzymes are really not JH-directed, but will, under experimental conditions, generate metabolites not seen under in vivo conditions. Analysis of JHE activity is further complicated by the existence of both hemolymph and tissue JH binding proteins that preferentially bind certain homologs (Goodman et al., 1976; Park et al., 1993) and enantiomers (Schooley et al., 1978b). These proteins have a significant influence on JHE activity (Hammock et al., 1975) due to preferential binding of one enantiomer (Peter, 1990) or homolog (Halarnkar et al., 1993) over another (see Section 3.7.8.2.1). The recent development of chromatographic methods to separate racemic preparations of JH should alleviate some of these problems (Cusson et al., 1997). Finally, in the drive to discover a unifying theory to explain JH catabolism, the diversity of the class Insecta is often overlooked. While synthesis of the results of different studies is unavoidable, to assume that all insect species use a single common pathway for JH catabolism would be a gross

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oversimplification of the real situation. One need only compare JH metabolism in the flea Ctenocephalides felis, which utilizes JHE (Keiser et al., 2002), D. melanogaster, which utilizes JHEH (Casas et al., 1991), and the moth T. ni, which utilizes both pathways (Kallapur et al., 1996), to understand that multiple pathways for JH catabolism have evolved in this diverse class.

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The role of JHs in embryonic development remains an enigma despite the discovery of the hormones in the eggs of H. cecropia more than 40 years ago (Gilbert and Schneiderman, 1961). However, the paradigm that so elegantly describes the developmental action of JH during larval to pupal metamorphosis has not, until recently, seemed applicable to the embryonic development of insects. The observation that JH titers rise at different times during embryogenesis of hemimetabolous and holometabolous insects has led Truman and Riddiford (1999) to speculate that JH may have played a significant role in the evolutionary divergence of these two groups. As the evidence presented below indicates, the role of JH during embryonic development warrants further consideration. 3.7.10.1. Juvenile Hormone Homologs and Precursors Present during Embryogenesis

The JH homologs from embryos of only a few species have been identified thus far (Table 1), giving us a very incomplete picture. What does emerge from these limited data is confirmation that JH III in nonlepidopteran insects is the predominant

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embryonic hormone, and following the pattern observed in other stages (Schooley et al., 1984). Furthermore, JH homologs and precursors not detected in larvae or adults exist in eggs, including JH 0, 4-methyl JH I, and the JH precursor, MF. Even more remarkable are the measurable titers of these homologs and precursors in whole-body extracts of embryos. While it is difficult to compare titers in whole-body extracts with those in hemolymph, the levels of JH in the embryo do appear significantly higher than those in hemolymph at any time during larval development (Edwards et al., 2001). 3.7.10.2. Role of Methyl Farnesoate during Embryogenesis

One of the more interesting JH-like molecules found in embryos is MF, the final precursor in the pathway to JH III biosynthesis. While MF possesses the farnesyl backbone of the JHs, it lacks the terminal epoxide at the C10 position (Figure 1) and exhibits relatively low activity in bioassays when compared to JH III (Sla´ ma et al., 1974). Several investigators have reported high levels of JH III and MF in N. cinerea midway through embryonic development (Baker et al., 1984; Lanzrein et al., 1984; Bru¨ ning et al., 1985). Using the RCA, Bu¨ rgin and Lanzrein (1988) demonstrated that the embryonic CA are indeed synthesizing JH III and MF. The CA of N. cinerea become differentiated just before dorsal closure of the embryo, and synthesis of MF and JH III begins shortly thereafter (Bu¨ rgin and Lanzrein, 1988). MF levels rise first, followed by JH III about a day later. Both MF and JH III levels remain elevated for several days, exceeding 800 ng g1, and then decline during the later stages of embryonic development with MF decreasing to undetectable levels (Bru¨ ning et al., 1985).

Table 1 Juvenile hormone homologs present during insect embryogenesis Species

Hormones

Reference

Thermobia domestica Blatella orientalis Nauphoeta cinerea Locusta migratoria Telogryllus commodus Oncopeltus fasciatus Leptinotarsa decemlineata Melolontha melolontha Heliothis virescens Spodoptera littoralis Manduca sexta Hyalophora cecropia Bombyx mori Apis mellifera

JH III JH III JH III, methyl farnesoate JH III JH III JH III JH III JH III JH 0, I, II JH I, II, III JH 0, I, II, 4-Me JH I JH 0, I, II JH I, II, III JH III

Baker et al. (1984) Short and Edwards (1992) Baker et al. (1984), Bru¨ning et al. (1985) Temin et al. (1986), Pener et al. (1986) Loher et al. (1983) Bergot et al. (1981a) deKort et al. (1982) Trautmann et al. (1974) Bergot et al. (1981a) Steiner et al. (1999) Bergot et al. (1981a) Bergot et al. (1981a) Gharib et al. (1983) Rembold et al. (1992)

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Recent studies on another cockroach, D. punctata, support the hypothesis that MF is a major product of the embryonic CA (Stay et al., 2002). Using the RCA to monitor rates of MF and JH III synthesis, these investigators demonstrated that production of MF increases approximately a day before that of JH III and, as in the case of N. cinerea, synthesis of both compounds begin shortly after dorsal closure. Ultrastructural studies of the CA confirm the synthetic activity of the glands after dorsal closure (Lee and Chiang, 1997). In contrast to N. cinerea, where MF and JH III are found in equivalent amounts, the biosynthetic rate of JH in D. punctata exceeds that of MF by 30-fold. Stay et al. (2002), examining the role of D. punctata allatostatin 7 (see Section 3.7.7.2) in regulating MF and JH biosynthesis in D. punctata embryos, found the process complex. This potent inhibitor of JH III biosynthesis in adults also inhibits synthesis of MF and JH after dorsal closure. However, prior to dorsal closure, the allatostatin has the reverse effect, stimulating MF biosynthesis. Prior to innervation by neurons transporting the allatostatin, the CA produce MF and can be stimulated by DippuAST to produce MF in a dose-dependent manner. Following innervation by Dippu-AST-containing neurons, the glands begin to synthesize both MF and JH, and exogenous Dippu-AST downregulates synthesis of both molecules. A study with N. cinerea embryos is a reminder that MF and JH III cannot be viewed as equivalent or interchangeable (Bru¨ ning and Lanzrein, 1987). Treatment of N. cinerea embryos with ethoxy-precocene eliminates JH III titers but has little effect on MF levels; yet, the investigators were able to rescue ethoxy-precocene treated embryos with JH III. Interestingly, although ethoxy-precocene is known to induce destruction of the CA in certain insect species, it does not chemically allatectomize this species. Histological studies revealed that as late as 10 days after ethoxy-precocene treatment, the CA were similar to control CA in size and cell number but the nuclei appeared pycnotic. It remains to be seen whether MF is important to embryonic development in N. cinerea and D. punctata, or in any other species. 3.7.10.3. Juvenile Hormone Titers during Embryogenesis: Correlation with Developmental Events

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As noted previously (see Section 3.7.10.1), the action of JH during embryogenesis is unclear and may be different from its role of maintaining the status quo role during the larval stage. Associating changes in JH titers to events in organogenesis could provide

clues to the role(s) of JH during embryogenesis. Short and Edwards (1992) measured JH titers in embryos of the cockroach Blattella orientalis, and demonstrated that JH III is present at very low levels at oviposition and in the period preceding dorsal closure. Midway through embryogenesis, JH III titers rise dramatically, reaching better than 600 ng g1 of tissue before decreasing. Unfortunately, the investigators did not associate the JH titers with specific embryological events, thus making it difficult to link the peak with any specific developmental process. By contrast, Bru¨ ning et al. (1985) were able to demonstrate that JH III appears in the embryo of N. cinerea only after dorsal closure; no hormone was detected prior to dorsal closure. As in the case of B. orientalis, JH III in embryos of N. cinerea rises to a very high level (800 ng g1 of tissue). The titer remains high for about 8 days and then falls to undetectable levels prior to emergence. Temin et al. (1986) conducted a closely timed study of JH titers during embryonic development of L. migratoria. During the first 60 h after oviposition, JH III levels remain reasonably constant at about 15 pg per egg. Blastokinesis, a movement of the embryo in relation to the yolk mass that consists of two phases (anatrepsis and katatrepsis) (Johannsen and Butt, 1941), occurs from 60 to 108 h after oviposition. During this time, JH is undetectable; however, at dorsal closure, which occurs at approximately 120 h, JH titers begin to rise and reach their highest level at 180 h. Unlike the massive peak in the JH titer observed in B. orientalis and N. cinerea, the peak titer in L. migratoria is approximately 45–50 pg per egg. Levels then fall slowly and are undetectable at the time of nymphal emergence. It is interesting that no JH is detected during the period between the initiation of blastokinesis and dorsal closure, a time that corresponds to the most active period of organogenesis (Temin et al., 1986). However, as noted below, exposure to JH during this period can have profound effects on the development of the embryo and larva. Dorn (1975), using a bioassay to titer embryonic JH in Oncopeltus fasciatus, found that newly oviposited eggs contain low, but detectable, levels of JH. He suggests that the hormone found at oviposition is maternal in origin. The titers remain stable during the first 2 days of embryonic development, but then rise rapidly during the third day. This period in O. fasciatus is marked by the completion of CA development and katatrepsis. Levels of JH continue to rise during the fourth day of embryonic development, a period in which final dorsal closure occurs, and peak during the fifth day at levels nearly

p0945

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p0960

p9015

p0970

15 times that found at oviposition. JH levels drop at the time of hatching. Titers of JH in the embryos of Lepidoptera present a more confusing picture. Using a bioassay, Gilbert and Schneiderman (1961) were the first to demonstrate that unfertilized eggs of H. cecropia contained JH. Bergot et al. (1981a) subsequently used GC-MS to detect low levels of JH I at 108 h after fertilization, a period following dorsal closure in this species (Riddiford, 1970). After dorsal closure, JH titers rise to a peak at 156 h (520 pg tissue g1) and then slowly decline to 50 pg tissue g1 at the time of emergence, approximately 250 h after oviposition. In M. sexta embryos, organogenesis has been extensively studied and correlated to JH titers. JH titers are undetectable by GC-MS prior to provisional dorsal closure (54 h after oviposition) (Bergot et al., 1981a; Dorn et al., 1987). The titers rise to their highest level between 60 and 72 h after oviposition, when a true larval cuticle appears. Interestingly, this peak in JH is composed of several homologs, including JH 0 and 4-methyl JH I, that are unique to the embryonic stage of Lepidoptera. The titers drop to less than 50 pg g1 total JH at the time of hatching, approximately 120 h after oviposition. Using an HPLC/bioassay method to measure JH levels in the embryo of the noctuid T. ni, Grossniklaus-Bu¨ rgin and Lanzrein (1990) showed that JH titers rise during the period of early segmentation, prior to dorsal closure, and remain detectable throughout the period of early organogenesis. As has been observed in other species, the JH titer increases to its highest level shortly after dorsal closure and then declines at hatching. The extensive embryological landmarks available for T. ni permit a direct correlation of the peak in the JH titer with the formation of the first larval cuticle. A similar JH profile is observed in another noctuid, S. littoralis (Steiner et al., 1999). GC-MS quantitation was used to demonstrate that low levels of JH II are detectable prior to dorsal closure, during a period in which segmentation and limb growth is extensive. JH titers are at their highest after dorsal closure and more precisely, highest at the time the true larval cuticle is being formed. Titers drop as the embryo develops, but in contrast to other species where the hormone declines to undetectable levels at hatching, S. littoralis maintains elevated titers of JH through to emergence. Despite the extensive body of work concerning JH and caste differentiation in A. mellifera (Huang et al., 1991; Schulz et al., 2002) (see Chapter 3.13), there is little information linking embryological markers with JH titers in this species. Rembold

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et al. (1992) found that JH III levels are undetectable by GC-MS until very late in embryogenic development (66 h after oviposition). The late peak in the JH titer (72–76 h) may reflect the fact that the CA are not fully developed until 62–64 h after oviposition (Nelson, 1915). Moreover, dorsal closure occurs late in this species, (62–66 h) which is in keeping with the established developmental correlation between dorsal closure and the peak in the embryonic JH titer. It should be noted that the A. mellifera embryo does not undergo blastokinesis, thus making it difficult to relate changes in the JH titer with events surrounding and within this process. As previously noted, the very early presence of JH in the egg of some species has led several investigators to suggest the origin of the hormone at this stage is maternal. In other words, hemolymph JH from the adult female is sequestered by the embryo during the process of vitellogenesis. Gilbert and Schneiderman (1961), in their pioneering work on JH, demonstrated that newly oviposited eggs contain low levels of JH; however, allatectomy of adult gravid females leads to eggs devoid of JH. They, as well as later investigators (Temin et al., 1986; Steiner et al., 1999), proposed that JH is taken up passively by the embryo. Indeed, presupposing that all JH is derived from the CA, this premise would be true, but the possibility remains that extra-allatal sites are involved in JH biosynthesis. Hartmann et al. (1987) demonstrated that the serosa of L. migratoria is capable of methylation of JH III acid to form JH III. Whether this process can yield sufficient JH to account for the amount of JH detected before dorsal closure is uncertain, nor is it clear whether other insects utilize this unique tissue for biosynthesis of JH. A summary of JH titers during embryogenesis is seen in Table 2. Do these profiles indicate that JH is requisite at key times during embryonic development? Unfortunately, the markers that denote embryological development vary widely among species and make comparisons difficult. For example, blastokinesisis varies radically among species (Johannsen and Butt, 1941) and does not provide an adequate morphological marker to make generalizations. Moreover, while the movements of blastokinesis may appear similar in many species, this process may not necessarily signify a developmentally defining homologous event (Heming, 2003). By contrast, dorsal closure is an anatomical reference point that signifies a specific developmental event. However, an embryonic cell layer, more commonly referred to as a membrane, is known to create a provisional dorsal closure that occurs earlier than the final

370 The Juvenile Hormones

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Table 2 JH levels during insect embryogenesisa

Oviposition

Blastokinesis

Dorsal closure or larval cuticulogenesis

Lowb

None

Highd

Temin et al. (1986)

None Low

None Low

High High

Bru¨ning et al. (1985) Short and Edwards (1992)

Low

Mediumc

High

Dorn (1975)

Manduca sexta Spodoptera littoralis Trichoplusia ni

None Low None

None Not determined Low

High High High

Hyalophora cecropia

Low

Low

High

Bergot et al. (1981a) Steiner et al. (1999) Grossniklaus-Bu¨rgin and Lanzrein (1990) Gilbert and Schneiderman (1961), Bergot et al. (1981)

None

Nonee

High

Rembold et al. (1992)

None

Nonee

None

Bownes and Rembold (1987)

Order and species

Reference

Hemimetabolous

Orthoptera Locusta migratoria

Dictyoptera Nauphoeta cinerea Blatta orientalis

Hemiptera Oncopeltus fasciatus Holometabolous

Lepidoptera

Hymenoptera Apis mellifera

Diptera Drosophila melanogaster a

JH titers were determined by GC/MS or HPLC/bioassay. Low JH titers are those that are less than one-third of the highest level reported. c Medium JH levels are those that fall between one-third and two-thirds of the highest level reported. d High JH levels are those that are greater than two-thirds of the highest level reported. e Blastokinesis is not observed in these species; in these species, JH titers are reported for the period corresponding to approximately half way between oviposition and dorsal closure. b

dorsal closure. An example of the confusion this can cause is readily seen in studies of the embryo of M. sexta, where a provisional dorsal closure is evident at about 50 h after oviposition (Broadie et al., 1991), while the final dorsal closure becomes apparent only at 102 h after oviposition (Dorn et al., 1987). While knowledge of JH titers offers correlative information when linked to a key point in embryogenesis, it can provide only indirect evidence for the role(s) the hormone plays during this period. 3.7.10.4. Roles of Juvenile Hormone during Embryogenesis p0985

To better understand the embryological roles of JH, two experimental approaches have been taken: chemical allatectomy using precocene and the application of exogenous JH or a JH agonist. Application of JH I or II to eggs of Thermobia domestica, an ametabolous insect, any time from oviposition to 2 days before hatching, leads to highly deformed embryos and poor emergence rates (Rohdendorf and Sehnal, 1973). Injeyan et al. (1979) applied JH III to S. gregaria embryos and found a different pattern of JH-induced disruption. JH applied shortly after oviposition has little effect on embryogenesis, but when applied from

days 3 to 9 postoviposition has a profound effect on development, including disruption of blastokinesis, inhibition of postblastokinesis development, and failure of the vermiform larva either to initiate or to complete ecdysis. JH has no effect on embryonic development when applied between day 10 and day 16 (hatching). Disruption of development is accompanied by increased pigmentation of the embryos, and it was discovered that the pigmented regions of treated insects have a much thinner procuticle. In contrast to these results, SbrennaMicciarelli (1977) demonstrated that high doses of the analog farnesyl methyl ether applied to the embryo just before blastokinesis induces premature development of the cuticle. Although the results are conflicting, both authors conclude that JH may have a role in cuticulogenesis. Application of precocene to N. cinerea embryos following dorsal closure results in abnormal midgut development, which appears to stem from the failure of ectodermal cells to migrate from the developing stomodaeum and proctodaeum into the midgut rudiments to form the gut epithelium (Bru¨ ning et al., 1985). In addition, the absence of JH after dorsal closure leads to disintegration of the fat body and abnormalities in the cuticle. Oncopeltus fasciatus

The Juvenile Hormones

embryos treated with precocene at blastokinesis fail to undergo final dorsal closure, suggesting that JH is involved in development of ectodermal tissue (Dorn, 1982). This developmental aberration can be rescued by topical application of JH. Enslee and Riddiford (1977) demonstrated that blastokinesis in P. apterus was vulnerable to exogenous JH, which interfered with dorsal closure and induced truncated appendage development and early pigmentation of developing eye and abdominal structures. These authors suggested that the extraembryonic ‘‘membranes’’ might be the target of JH. Day-old embryos of R. prolixus, when treated with high levels of fenoxycarb, do not develop past katatrepsis, and patterns of protein synthesis, as determined by two-dimensional gel electrophoresis, are significantly altered (Kelly and Huebner, 1987). In yet another hemipteran, Aphis fabae, precocene inhibits embryogenesis, but the process can be rescued by JH (Hardie, 1987). Interestingly, JH appears to accelerate embryonic development via a mechanism independent of the parthenogenic reproductive strategy employed by aphids. The effect of JH or analogs on the embryogenesis of holometabolous insects is similar to that seen in hemimetabolous insects. Unfortunately fewer studies have been carried out and all of them utilize the addition of exogenous JH to determine an effect. In Lepidoptera, JH injected into the female prior to oviposition halts embryogenesis at formation of the blastoderm, while JH applied directly to the egg shortly after oviposition has no effect on blastoderm formation, but blocks blastokinesis (Riddiford, 1970). Even without blastokinesis, embryos continue larval differentiation and form recognizable but incomplete first instars, as in P. apterus (Enslee and Riddiford, 1977). In another lepidopteran embryo, that of Plodia interpunctella, JH agonists induce aberrant dorsal closure and a loss of trachea (Dyby and Silhacek, 1997). A very different situation exists in D. melanogaster, a species in which no JH is detected during embryogenesis (Bownes and Rembold, 1987). While JH analogs can disrupt development if applied very early, large doses of JH I do not have an effect on embryogenesis (Smith and Arking, 1975). The underlying molecular role of JH during embryogenesis is even more obscure. In an early study, Rao and Krishnakumaran (1974) attempted to elucidate the role of JH at the molecular level. When these investigators examined morphological alterations and changes in DNA synthesis in embryos of A. domesticus challenged with JH I, they found a qualitative drop in incorporation of thymidine into DNA, as monitored by autoradiography.

371

Embryonic epidermal tissue is the first to show reduced incorporation of thymidine in the presence of exogenous JH; however, some tissues appear refractory to the hormone. Interestingly, JH-treated A. domesticus embryos develop legs and antennae that are short and stumpy, which led Rao and Krishnakumaran (1974) to suggest that JH may suppress DNA synthesis and cell division in the embryonic appendages. Despite the paucity of information, it is tempting to speculate that embryonic ectodermal tissue is a major target of JH. The effects of JH on cuticular morphology, dorsal closure, and pigmentation of embryonic epidermal structures support this idea, and are in keeping with the function of JH during larval development. The fact that JH acts on midgut and fat body development in at least some species indicates there also may be a wider role for the hormone. 3.7.10.5. Juvenile Hormone Binding Proteins of the Embryo

The JHBPs have long been known to modulate the catabolism of hemolymph JH during the larval period (Hammock et al., 1975; Roe and Venkatesh, 1990; deKort and Granger, 1996) (see Section 3.7.9). It is generally assumed that JHBP has a unique hormone-binding domain whose steric arrangement hinders the access of catabolic enzymes to JH (Goodman, 1990). While the roles of the hJHBP in larval development are well defined (see Section 3.7.8.3), the role of embryonic JHBP is less clear. It is now clear that significant levels of JHBP are present throughout embryogenesis even though the circulatory system is not yet fully functional until late in development, and JH levels, as noted above, do not rise until the insect is midway through embryonic development. In L. migratoria, a JHBP similar to that found in nymphal hemolymph is present in both the embryo and serosa (Hartmann et al., 1987), prior to the development of the CA and a functional circulatory system. These investigators demonstrated that the serosa has a methyl transferase capable of converting JH III acid to JH III, and surmised that the embryonic JHBP serves either to distribute the hormone or to act as a buffer, protecting the embryo from excess maternal JH. The hypothesis of a protective function for embryonic JHBP is supported by the fact that JH levels in the gravid female are at least an order of magnitude higher than in the eggs (Temin et al., 1986). Thus, the JHBP could act as a ‘‘sponge’’, keeping the JH accessible to catabolic enzymes that can readily inactivate the hormone.

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372 The Juvenile Hormones

In M. sexta, a hJHBP-like molecule is present at oviposition and remains at detectable levels throughout embryogenesis (Touhara and Prestwich, 1994; Touhara et al., 1994). A re-examination of the characteristics and titers of the embryonic JHBP demonstrated that the embryonic JHBP is identical in binding characteristics (Touhara and Prestwich, 1994; Touhara et al., 1994), molecular mass, and immunochemical properties to the larval hemolymph form (Orth et al., 2003a). Furthermore, the nucleotide sequence of the embryonic JHBP gene is identical to that of the larval gene (Orth et al., 2003b); thus, it can be concluded that the embryonic JHBP is identical to hJHBP. Expression of the JHBP gene begins in both the embryo and the serosa approximately 15 h after oviposition, peaks between 24 and 36 h, and then declines to undetectable levels at 72 h. The burst of JHBP expression occurs before emergence of functional CA, fat body, or circulatory system. In contrast to its gene expression, titers of JHBP are at their highest during the first half of embryogenesis (Figure 17). The titer declines rapidly between 36 and 60 h after oviposition and then slowly drops to its lowest level at the time of emergence. The peak of gene expression does not appear to enhance the already high levels of JHBP and the reason for this temporal discordance between gene expression and the JHBP titer is unclear. As in the

case of L. migratoria, it appears that both the serosa and the embryo of M. sexta are capable of expressing JHBP. Curiously, the expression of JHBP in M. sexta begins about 15 h after oviposition, corresponding to the time at which the serosa becomes active in secretion of the first of several embryonic ‘‘membranes’’ (Lamer and Dorn, 2001). Most of the embryonic JHBP in M. sexta, especially in the early hours following oviposition, is maternally derived and is assumed to be sequestered from hemolymph during the process of vitellogenesis. The presence of relatively high levels of the protein prior to development of the CA or circulatory system suggests that, as in L. migratoria, high levels of JHBP in the early embryo of M. sexta ensure low levels of unbound JH, thus protecting the embryo from excess JH (Orth et al., 2003b). JH bioassay data indicate that hemolymph JH titers in adult female M. sexta are as high as those found in the early fourth stadium larvae (Judy, personal communication). Thus, during the early part of embryogenesis, the maternally derived JHBP acts as an effective buffer, constantly binding and releasing the hormone in accordance with equilibrium conditions. Embryonic JHE displays a developmental profile similar to that of JHBP (Share et al., 1988) and together, these proteins regulate bioavailability of the hormone to the developing embryo.

Figure 17 Titers of JH and juvenile hormone binding protein during embryological development of M. sexta. Bars, JHBP mean titers (
SD) (JHBP adapted from Orth, A.P., Tauchman, S.J., Doll, S.C., Goodman, W.G., 2003b. Embryonic expression of juvenile hormone binding protein and its relationship to the toxic effects of juvenile hormone in Manduca sexta. Insect Biochem. Mol. Biol. 33, 1275–1284. Line (—), total JH titer data adapted from Bergot, B.J., Baker, F.C., Cerf, D.C., Jamieson, G., Schooley, D.A., 1981a. Qualitative and quantitative aspects of juvenile hormone titers in developing embryos of several insect species: discovery of a new JH-like substance extracted from eggs of Manduca sexta. In: Pratt, G.E., Brooks, G.T. (Eds.), Juvenile Hormone Biochemistry. Elsevier/North-Holland, Amsterdam, pp. 33–45.

3.7.10.5.1. Juvenile hormone catabolism during embryogenesis: methyl ester hydrolysis Embryonic titers of JH are the result of both biosynthesis and catabolism. While this area has received considerable attention in the larval stage (see Section 3.7.9), scant attention has been focused on these catabolic enzymes in the egg. Roe et al. (1987a, 1987b) examined the metabolism of JH in the eggs of the cricket A. domesticus. Their studies found that the major route of metabolism is the conversion of JH to JH acid. They reported that general esterase activity, as determined using a-naphthyl acetate as a substrate, remains relatively constant throughout embryonic development. In contrast, JHE activity is high in unfertilized eggs and remains high until the time of dorsal closure, when it drops to its lowest level and remains relatively low until nymphal emergence. In a very thorough analysis of JH titers (Bru¨ ning et al., 1985), JH biosynthetic rates, and JHE activity in N. cinerea embryos, Lanzrein and her colleagues demonstrated that JHE activity is very low following dorsal closure (20 days after oviposition) (Bu¨ rgin and Lanzrein, 1988). Figure 18 shows that the massive peak representing both JH III and MF begins to decline very rapidly when JHE activity

The Juvenile Hormones

373

Figure 18 JH titers, JH biosynthetic rates, and JHE activity from Nauphoeta cinerea embryos. Closed circles, JH III titers (Bru¨ning et al., 1985); bars, relative JH III biosynthesis as determined by the radiochemical assay (Bu¨rgin and Lanzrein, 1988); closed triangles, JHE activity (Bu¨rgin and Lanzrein, 1988). Dorsal closure occurs at day 20.

p1040

rises at day 35. According to these investigators, the rise in JHE corresponds to the period of chorion breakdown; as a result, the increase in available oxygen significantly changes the physiology of the rapidly developing embryo. Interestingly, JHE activity does not appear to be localized to the hemolymph, but rather is distributed generally throughout the entire embryo (Bu¨ rgin and Lanzrein, 1988). Since JH III titers in the first instar nymphs are low (1ng g1), the dramatic rise in JHE activity may be necessary to clear both JH and MF prior to nymphal emergence (Bru¨ ning et al., 1985). The rapid rise in JHE activity observed at the end of embryonic development in N. cinerea contrasts with that observed in M. sexta. In this insect, JHE activity is high in preovipositional eggs, but declines well before provisional dorsal closure (Share et al., 1988). JHE activity remains relatively constant throughout the rest of embryonic development and does not show the dramatic increase in activity that is observed in N. cinerea (Bu¨ rgin and Lanzrein, 1988). Despite the presence of JHE activity, JH titers increase dramatically in the period surrounding larval cuticulogenesis (Bergot et al., 1981a). The rise in the JH titer, in the presence of significant JHE activity, contradicts the generally held view that JHE is involved in inactivation of the hormone (see Section 3.7.9.1). It may well be that JH and JHE are sufficiently compartmentalized at the onset of cuticulogenesis that catabolism of the hormone is limited. Once the larval circulatory system is fully functional, approximately 24 h later as determined by vigorous dorsal vessel contraction (Broadie et al., 1991), contact between the hormone and the catabolic enzymes result in a reduced JH titer as the enzyme carries out its anticipated function.

3.7.10.5.2. Juvenile hormone catabolism during embryogenesis: epoxide hydrolysis While most studies on JH catabolism have been focused on the JHEs, a handful of investigators have examined the action of the JHEH (see Section 3.7.9.2). Share et al. (1988), in their studies on JH metabolism during embryogenesis in M. sexta, demonstrated that JHEH activity is responsible for only a small amount of the total catabolic activity. They found that this enzyme hydrolyzes JH III about six times better than JH I, but they did not take into account the fact that JH I binds to the JHBP with a significantly higher affinity and thus protects the hormone from degradation (Park et al., 1993). The first characterization of a specific JHEH early in development was carried out by Touhara and Prestwich (1993) in a study of JH metabolism in eggs of M. sexta. Using the JH photoaffinity label [3H]epoxyfarnesyl diazoacetate, these investigators demonstrated that the radiolabeled analog covalently attaches to two proteins in the egg, the juvenile hormone binding protein and a 50 kDa protein that was identified as a JHEH. The majority of the JHEH activity is found in the microsomal fraction of the egg homogenate. The purified JHEH displays an apparent Km of 0.61 mM for JH I and 0.28 mM for JH III, confirming the earlier observation by Share et al. (1988) that JH III appears to be the preferred substrate. A subsequent characterization of recombinant JHEH from M. sexta reconfirmed this substrate specificity (Debernard et al., 1998). This study also found that the hJHBP protects the hormone from epoxide hydration, but whether this occurs in vivo is uncertain since the binding protein is present in the hemolymph (with the exception of the fat body), while the JHEH is localized to the cellular compartment.

374 The Juvenile Hormones

3.7.10.6. Juvenile Hormones and the Evolution of Insect Metamorphosis

Truman and Riddiford (1999, 2002) have proposed that the temporal appearance of JH during embryonic development may have played an important role in the evolution of holometabolic insects. Their hypothesis is based, in part, on the idea that ancestral insects displayed three stages after embryogenesis: the pronymph, the nymph, and the adult. The pronymphal stage represents a specialized hatching stage in ametabolous and hemimetabolous insects that is (1) nonfeeding, (2) often mobile, (3) often surrounded by an embryonic cuticle, and (4) different in body proportions from that of the first instar. Depending upon the species, the pronymphal stage is initiated after completion of blastokinesis and may end prior to hatching or persist for several days after hatching. The Truman and Riddiford hypothesis states that the pronymphal and nymphal stages of this insect are developmentally equivalent to the larval and pupal stages, respectively, of the holometabolus insect. Moreover, there is a requirement for a JH-free period in ametabolous and hemimetabolous species that allows nymphal development to proceed towards adult stage. Can experimental manipulation mimic the evolutionary processes that led to the transition to the holometabolous state? As noted earlier, JH can induce certain tissues in hemimetabolous species to undergo apparent premature differentiation before those tissues are completely patterned. This curious phenomenon has led Truman and Riddiford (1999, 2002) to suggest that the early appearance of JH during embryogenesis, either experimentally or naturally, modifies the developmental trajectory of the holometabolous insect to the adult stage, giving rise to a protolarval stage. From an evolutionary standpoint, advancing the time at which JH appears during embryogenesis may have been crucial to the transition from the pronymphal stage of the hemimetabolous insect to the protolarval stage of holometabolous species. While this hypothesis deserves much merit for simplicity and elegance, it is not without criticism (Heming, 2003). As demonstrated in Table 2, JH titers have been rigorously determined in only a handful of species, thus leaving any generalizations about the endocrine-driven rise of holometabolous development open to question. Moreover, working definitions of key embryogenic events, such as blastokinesis and dorsal closure, may be interpreted differently, depending on the investigator. The problem is further compounded by contradictory data on the time at which JH titers rise in relation to the key embryonic events. Truman and

Riddiford (2002) suggest that JH titers in the embryo of the lepidopteran M. sexta are highest at the time of katatrepsis, yet Ziese and Dorn (2003) place katatrepsis at a time when JH titers (Bergot et al., 1981a) are undetectable. Assigning an embryological role to JH when the levels are low raises yet another issue: low JH titers at oviposition and at blastokinesis in some species contradict the hypothesis that a JH-free period is required for further normal development (Table 2). While the Truman and Riddiford hypothesis about the rise of holometabolous insects presents an intriguing model that deserves further critical study, the role that JH may have had in this process is still open.

3.7.11. Juvenile Hormones in Premetamorphic Development 3.7.11.1. Juvenile Hormone Titers

The premetamorphic titers of JH appear to vary greatly among the insect orders, with some orders, such as Dictyoptera, displaying levels 100-fold greater than Lepidoptera (Gilbert et al., 2000). Yet one pattern has remained constant since the discovery of JH in the 1930s: JH titers, on average, are high while the larva is growing and feeding but drop at a well-defined point to permit metamorphosis. This pattern can be further refined to distinguish between hemimetabolous and holometabolous insects (Riddiford, 1994). In hemimetabolous insects, JH titers are low to undetectable during the final stadium. In holometabolous insects, JH titers are relatively high at the beginning of the final larval stadium but decline to undetectable levels prior to the cessation of feeding. The absence of JH permits the release of the prothoracicotropic hormone (PTTH) from specific cerebral neurosecretory cells (Rountree and Bollenbacher, 1986), and PTTH then induces ecdysteroid synthesis by the prothoracic glands, initiating a small rise in hemolymph ecdysteroid levels (Bollenbacher et al., 1981) (see Chapter 3.2). This rise in the ecdysteroid titer in the absence of JH initiates metamorphosis. In contrast to the hemimetabolous insects, a second increase in the hemolymph JH titer is typically observed after the insect has found a suitable pupation site. This peak in the titer is thought to prevent precocious adult differentiation of imaginal discs and other imaginal precursors (Riddiford, 1994). 3.7.11.2. Potential Problems with Titer Determinations

One of the major impediments to better understanding the role(s) of JH is the lack of precise titers

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during biologically relevant periods. Virtually all hormones studied in vertebrates display daily oscillations in their titer and release and are, in many cases, linked to critical homeostatic events (Czeisler and Klerman, 1999). Early studies demonstrated a daily, bimodal, rhythmic fluctuation in the size of the nuclei in CA of adult D. melanogaster (Rensing, 1964). While size of the CA cells does not necessarily indicate a gland active in JH biosynthesis (Tobe and Stay, 1985), there is ample evidence that indicates enlarged glands are biosynthetically active (Chiang et al., 1989, 1991). Thus, fluctuations in CA cell size suggests circadian fluctuations in JH synthesis. The importance of examining titer fluctuations in detail can be seen in the elegant work from Steel’s laboratory, which demonstrates a circadian rhythmicity of ecdysteroid titers in nymphal R. prolixus (Steel and Ampleford, 1984; Vafopoulou and Steel, 2001) (see Chapter 3.11). A single-point JH determination every 24 h may not be a serious problem for the student of metamorphosis, where JHregulated events stretch over a period of several days. However, it does make a difference when relatively rapid changes in JH titer regulate critical cellular events that occur during a short time period. Recent studies demonstrating that JH titers fluctuate rapidly in adult insects supports the premise that detailed studies of JH titers during the premetamorphic stages are needed. Elekonich et al. (2001) found that honeybee foragers show a diurnal increase in hemolymph JH titers, from about 100 ng ml1 in the late morning to over 350 ng ml1 by late evening, a period spanning less than 12 h. A similar change is seen in the fourth stadium of M. sexta, where titers initially rise and then drop significantly over an 8 h period and may have some influence on levels of the transport protein, hJHBP (Fain and Riddiford, 1975; Orth et al., 1999). This is not to infer that JH titers must be absolutely accurate; the important information is knowledge of changes in the relative titers at well-defined times. However, even if precise staging is possible, the measurement of JH in samples from species or stages displaying low levels of hormone, or from smaller sized species, requires that tissue collections be pooled from a number of individuals. Valuable information about population variability is thus lost. Ideally, a single individual should be sampled sequentially over time, but this process leads to wound-induced changes (Caveney, 1970) that may significantly alter JH titers. In our hands, wounding has rapid and significant effects on hJHBP mRNA expression, which potentially modulates JH titers (Orth and Goodman, unpublished data). Even

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moderate handling prior to sample collection may have an effect on JH titers (Varjas et al., 1992). 3.7.11.3. Premetamorphic Roles of the Juvenile Hormones

The roles of JH in the stages preceding metamorphosis to the adult are ubiquitous, affecting behavior, organs and tissues, cellular organelles, and biochemical pathways. Several excellent reviews have covered these areas (Nijhout, 1994; Riddiford, 1994, 1996), and the focus here is on research since that time. 3.7.11.3.1. Behavioral and neuronal responses There are a number of behavioral phenomena ascribed to JH during the adult stage, including pheromone production and calling (review: Cusson et al., 1994), migration (review: Dingle and Winchell, 1997), phonotaxis (Stout et al., 1992, 1998; Bronsert et al., 2003), and caste determination and anatomical changes in the brain of honeybees (reviews: Robinson and Vargo, 1997; Elekonich and Robinson, 2000) (see Chapter 3.13). However, little is known about the effect of JH on behavior during the premetamorphic period, and most of these studies have been done in social or migratory insects. Late in the third stadium, honeybee larvae destined to become queens have JH titers five times higher than larvae destined to become workers (Rachinsky and Hartfelder, 1991; Rembold et al., 1992) (see Chapter 3.13). In another hymenopteran, the ant Phiedole bicarinata, large doses of methoprene, if applied during a critical period in the last stadium, induce worker-destined larvae to become soldiers (Wheeler and Nijhout, 1981). Higher JH titers during the last stadium of certain migratory species appear to elicit a stationary adult stage, rather than migration (Yagi and Kuramochi, 1976; Nijhout and Wheeler, 1982). Yin et al. (1987) demonstrated that methoprene can affect the circadian system in the lepidopteran Diatraea grandiosella, inducing a dose-dependent phase shift in adult eclosion. At the neuronal level, most endocrine studies have focused on the effect of ecdysteroids on the neural circuitry during metamorphsis (see Chapter 2.4), although JH appears to play a role as well. Truman and Reiss (1988) demonstrated that reorganization of neurons during metamorphosis of M. sexta is, in part, under the control of JH, but the regulatory elements involved are still undefined. Recent work suggests that JH affects larval neurons innervating the prothoracic gland of the cockroach P. americana (Richter and Gronert, 1999). Exposure of the insect to exogenous JH III and methoprene both in vivo and in vitro induces a short-term depression of spike

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activity in neurons innervating the prothoracic gland, but has no effect on the nervous connectives of the stomatogastric nervous system. This response to JH III is very rapid, occurring within 3 min of treatment and reaching a low (75% reduction) within 15 min. Curiously, fenoxycarb, a potent JH analog, was only half as active as JH III and methoprene; methyl laurate, a control lipid, had no effect. The authors speculate that JH is acting directly on membrane receptors to elicit the reduced neurotropic activity; this observation relates to an earlier hypothesis that JH can influence inhibitory neurotransmitter receptors such as g-aminobutyric acid (GABA) (Stout et al., 1992).

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3.7.11.3.2. Epidermal responses It has long been known that JH has a marked influence on epidermal and cuticular structure. The early work of Williams (1952) demonstrated that JH-containing extracts, when applied at a developmentally sensitive time during the pupal period, can induce a second pupal cuticle. This discovery prompted Williams to suggest that JH is a ‘‘status quo’’ hormone, a label that is still in frequent use (Willis, 1996). The integument of the insect is composed of an outer layer of secreted proteins, lipids, pigments, and complex carbohydrates termed the cuticle. The single layer of cells immediately beneath the cuticle, the epidermis, is responsible for synthesis and secretion of most, but not all, of the proteins found in the cuticle (Willis, 1996). The epidermis itself contains another subset of proteins, pigment-associating proteins, the expression and position of which in the cells appear to be regulated by JH. The radical changes in morphology at metamorphosis led Wigglesworth (1959) to predict that each stage of development had its own set of unique genes. While there does appear to be a limited group of stage-specific cuticular proteins, especially in the case of the higher flies, current evidence demonstrates that many cuticular proteins can be found in more than one stage of the life cycle (review: Willis, 1996). Hormonal regulation of epidermal gene expression has been examined in a number of insect species, but the most extensive studies, by Riddiford and her colleagues, have been carried out on the epidermal genes of M. sexta (Riddiford, 1994, 1996; Riddiford et al., 2001, 2003). These include the larval cuticular protein 14 (LCP14) (GenBank Accession no. 813279) (Rebers and Riddiford, 1988), LCP14.6 (GenBank Acc. No. U65902) (Rebers et al., 1997), LCP16/17 (GenBank Accession no. M25486) (Horodyski and Riddiford, 1989), the biliverdin-associating proteins, insecticyanin a and b (GenBank Accession nos. 864714

and 864715) (Li and Riddiford, 1992), and dopa decarboxylase (DDC) (GenBank Accession no. U03909) (Hiruma and Riddiford, 1988; Hiruma et al., 1995). Studies using the epidermis are particularly compelling, since the results of in vitro manipulations mirror those observed in vivo, and importantly, the genomic structure and flanking regions of their genes have been determined. LCP14 is a larval-specific gene that encodes for a 14 kDa protein expressed during the feeding period of each stadium (Riddiford, 1994). At the onset of a larval molt, mRNA for LCP14 rapidly becomes undetectable and remains so until the insect molts to the next stadium, at which time the expression levels rise again and the process repeats itself. In the last stadium, levels of LCP14 message rise during the first several days, but fall sharply prior to wandering and are no longer detectable. In vitro manipulation indicates that 20E suppresses the expression of this gene. If JH is present when 20E titers rise, suppression by 20E is only transient; if JH is absent, the gene is permanently silenced. While it is uncertain how JH acts at the molecular level to maintain the expression of LCP14, its function may involve the transcription factor bFTZ-F1. It has been shown that there are three potential binding sites for bFTZ-F1 in the LCP14 gene, one approximately 2 kb upstream from the start site and two in the first intron (Lan, personal communication). The expression of bFTZ-F1 begins about 16 h before molting, peaks at about 9 h before molting, and then ceases approximately 3 h before molting (Weller et al., 2001). Functional analysis of the putative LCP14 promoter indicates that the bFTZ-F1 response elements in the first intron are involved in downregulating LCP14 expression (Lan, personal communication). The decline in bFTZ-F1 expression coincides with the rise in JH at the very beginning of the fifth stadium and with it, the increase in LCP14 expression. Whether JH acts directly on the gene, or is involved in modulating the ecdysteroid effect, remains unclear. LCP14.6 is another hormonally controlled cuticular gene, which, like LCP14, is downregulated by 20E, but in contrast to LCP14, is suppressed in vitro by large doses of methoprene (Riddiford, 1986). Its expression is temporally and spatially complex and is not stage-specific, since it occurs in the larval, pupal, and adult stages. A comparison of its expression pattern with the JH titer profile suggests that, in vivo, LCP14.6 expression may be suppressed by the hormone. LCP16/17 encodes a multigene family of three proteins that appear midway through the feeding period of the fifth stadium (Horodyski and Riddiford, 1989). The developmental appearance of

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these proteins correlates with a thinning of cuticular lamellae and a corresponding increase in stiffness. Like LCP14.6, expression of the LCP16/17 is suppressed by large doses of methoprene. One of the more striking effects elicited by JH is its action on larval pigmentation (Nijhout, 1994; Applebaum et al., 1997). Nowhere is this more evident than in larval M. sexta, where two mutants have been discovered with anomalous pigmentation and aberrant JH titers. In the bl mutant (Safranek and Riddiford, 1975), JH titers are low at certain critical developmental points and the larvae are highly melanized; in the white mutant, the opposite is the case (Panchapakesan et al., 1994). To maintain the normal wild-type phenotype following a molt, i.e., a transparent cuticle and an epidermis with intracellular vesicles containing the biliverdinassociating protein, insecticyanin (Riley et al., 1984; Goodman et al., 1985), JH titers must be sufficiently high at the time of head capsule slippage (Hiruma and Riddiford, 1988; Riddiford, 1994). If JH titers are too high at the time of head capsule slippage, the resulting cuticle will be transparent, and the underlying epidermal cells will lack insecticyanin vesicles, causing the insect to appear white (Panchapakesan et al., 1994). Conversely, if JH titers are too low at the time of head capsule slippage, the resulting cuticle will be highly melanized and the epidermis devoid of insecticyanin vesicles, causing the insect to appear black (Goodman et al., 1987). Insecticyanin (Ins) is a 21 kDa protein that associates with biliverdin IX to yield an intensely blue protein important to larval camouflage (Goodman et al., 1985). Ins is found in epidermal cells, where it is sequestered in 1 mm vesicles (Figure 19). These vesicles, together with other epidermal and cuticular pigments, give the wild-type insect its distinctive blue–green hue during the feeding period (Riddiford et al., 1990). Analysis of epidermal ultrastructure indicates that the densely packed vesicles appear to be held in position by a web of microtubules (Goodman, unpublished data). A typical wild-type epidermal cell contains approximately 100 of the membrane-coated vesicles, localized in the apical region. The epidermal cell also secretes significant quantities of Ins into the hemolymph, making it one of the more prominent hemolymph proteins (Riddiford et al., 1990). At commitment to pupation, when JH is no longer present and ecdysteroid levels rise, the entire population of Ins-containing vesicles is secreted from the epidermal cell (Sedlak et al., 1983) and the insect ceases production of the protein (Goodman et al., 1987; Riddiford et al., 1990). Interestingly, the bl mutant, which has a low JH titer at head capsule slippage, lacks the

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Figure 19 Electron micrographs of the epidermis of the bl strain of M. sexta. Fourth instars, just prior to head capsule slippage, were treated with either acetone (a) or 50 ng JH I dissolved in acetone (b). Tissue was collected approximately 30 h later, at the time of the fifth larval ecdysis, and prepared for microscopy. C, cuticle; IV, insecticyanin vesicle; N, nucleus. Scale bars represent 1 mm. (Goodman, unpublished data.)

epidermal Ins-containing vesicles, thus mimicking the ultrastructure of the wandering fifth stadium epidermis. Topical application of JH to bl mutants, just prior to head capsule slippage, will induce the appearance of the Ins vesicles in the next stadium, similar in number and position to those in the wildtype (Goodman et al., 1987) (Figure 19). Moreover, JH, when applied at head capsule slippage, prevents cuticular melanization in the next stadium (Safranek and Riddiford, 1975). The role of JH in this process is still unclear but ultrastructural data suggests that the cytoskeleton may be involved in retaining the Ins vesicles in the apical portion of the epidermal cell. In addition to the presence of Ins mRNA in the epidermis of wild-type larvae, Ins mRNA has also been found in the fat body (Li and Riddiford, 1994; Li, 1996); however, the protein could not be detected immunologically unless pericardial cells were included in the preparation (Goodman et al., 1987). Curiously, JH appears to upregulate epidermal Ins mRNA abundance, while it downregulates this mRNA in the fat body (Li, 1996). As in the case of LCP14, 20E downregulates the epidermal Ins mRNA (Riddiford et al., 1990) during the molting period.

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Cloning of the Ins gene revealed a pair of duplicated genes termed Ins a and Ins b (Li and Riddiford, 1992) that are expressed in both the epidermis and fat body. Only the product encoded by the Ins b gene is found in the hemolymph (Riddiford, 1994). The difference in expression patterns between Ins a and Ins b led Li (1996) to investigate the structure of the genes, to determine whether different promoters were involved. Each of the genes displays a unique sequence in the 30 noncoding region, suggesting differential expression. Preliminary computational analysis of the 50 upstream region of Ins a indicated no known response elements (Lan, personal communication). DDC is the enzyme responsible for catalyzing the conversion of dopa to dopamine (Hiruma and Riddiford, 1984) while granular phenoloxidase (PO) catalyzes the oxidative dehydrogenation of diphenols to quinones (Hopkins and Kramer, 1992) (see Chapter 4.4). For cuticular melanization to occur, vesicles containing dopamine, the granular proenzyme form of PO, and other enzymes necessary for melanin synthesis are secreted into the new endocuticle following head capsule slippage. If JH titers are low or absent at the time of head capsule slippage, the proenzyme form of PO will be synthesized by the epidermis and deposited in the endocuticle for a period of approximately 12–14 h. Approximately 3 h before ecdysis, PO is activated, possibly by proteolytic activation (Jiang et al., 2003), and melanin begins to appear in the new cuticle. Topical application of JH to bl larvae, or to wild-type larvae that have been neck-ligated at head capsule slippage, will block phenoloxidase activity, and a new transparent cuticle will be formed (Riddiford et al., 2003). While JH suppresses cuticular melanization in a striking fashion, it remains unclear whether the hormone is acting on transcription or translational events that regulate PO activity. Hormonal regulation of epidermal DDC (EC 4.1.1.28) in D. melanogaster and M. sexta has been extensively studied by the Hodgett (Chen et al., 2002a, 2002b) and Riddiford laboratories (Riddiford, 1994; Riddiford et al., 2003). In M. sexta, 20E works directly on epidermis to block DDC synthesis; however, as the ecdysteroid titers decline during the head capsule slippage period, levels of DDC begin to rise and they peak near the time of the larval molt. A study of the 50 upstream region of the DDC gene indicates that there are several transcription factor binding sites, including a bFTZ-F1 response element between the TATA box and the transcriptional start site (Hiruma et al., 1995). Recently, Beckstead et al. (2001)

demonstrated that the D. melanogaster gene bonus (bon) encodes a homolog of the vertebrate TIF1 transcriptional cofactors. Among the many phenotypes associated with this gene is pigmentation (Beckstead et al., 2000). Bon binds to the AF-2 activation domain present in the ligand-binding domain of bFTZ-F1 and behaves as a transcriptional inhibitor in vivo. A transcription factor implicated in regulation of melanization as well as metamorphosis is the Broad-complex (Br-C). In the metamorphosis of D. melanogaster, Br-C is a required mediator in the DDC response to ecdysteroids. At pupariation and at eclosion, Br-C uncouples DDC from an active silencing mechanism that functions through two distinct cis-acting regions of the DDC locus (Chen et al., 2002a, 2002b). While much is known about DDC and its regulation via ecdysteroids, the role of JH in its control remains unclear (Hiruma et al., 1995; Chen et al., 2002a, 2002b; Riddiford et al., 2003). It has been demonstrated that the epidermis of allatectomized M. sexta larvae displays approximately 50% more DDC activity than epidermis from sham-operated insects, and that activity of the enzyme can be suppressed by moderate levels of JH I. While this observation is certainly provocative, it remains to be seen whether the JH effect on DDC expression is direct, or as some speculate, indirect (Hiruma, personal communication). 3.7.11.3.3. Fat body responses The premetamorphic fat body is responsible for metabolism of nutrients, synthesis of most hemolymph proteins, and detoxification of xenobiotics (Locke, 1984; Sondergaard, 1993; Haunerland and Shirk, 1995). Given the central roles this tissue plays, it is not surprising that an extensive body of literature has focused on the role of JH in regulating cellular and molecular events occurring in the fat body. Numerous studies involving hormonal regulation of ultrastructural changes in adult fat body, especially those regarding vitellogenesis (see Chapter 3.9), have been conducted, but few have focused on the ultrastructural changes induced by JH in the larval fat body. Such experiments are problematic since JH titers are already relatively high during most of larval life, and challenging the fat body with additional JH may result in pharmacological responses that mask the actual underlying events. Thus, the most comprehensive studies are developmental, correlating JH titers with ultrastructural changes occurring during the premetamorphic and metamorphic periods (Locke, 1984; Dean et al., 1985). In one series of studies, it was found that during the last stadium of both hemimetabolous

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and holometabolous insects, when JH titers are dropping, the fat body synthesizes and secretes several hexameric proteins collectively termed storage proteins (Levenbook, 1985). These proteins are retrieved from the hemolymph by the fat body and stored in relatively large vesicles for later use during the pupal–adult transformation (Locke, 1984; Dean et al., 1985). It has also been demonstrated that a JH analog affects fat body nuclei. Treatment of nymphal L. migratoria with high levels of methoprene leads to an increase in the size of fat body nuclei, which could be linked to either increased transcription rates or to increased ploidy levels (Cotton and Anstee, 1991). The latter option seems likely, since Jensen and Brasch (1985) demonstrated that allatectomy of adult L. migratoria prevents polyploidization, while methoprene restores the process. At the molecular level, the role of JH on fat body gene expression has been studied by a number of investigators in several different insect species. Certain of these gene targets are discussed below (see Section 3.7.12), and a comprehensive discussion of the various fat body genes thought to be under the control of JH has been presented by Riddiford (1994). One of the most well-studied groups of premetamorphic fat body genes and proteins is the hexamerins (Burmester et al., 1998; Burmester, 2002). The insect hexamerins encompass at least five different subgroups based on amino acid sequence, including: the coleopteran and dictyopteran JH-suppressible arylphorins, the lepidopteran JHsupressible hexamerins, lepidopteran aromatic amino acid-rich arylphorins, the lepidopteran methionine-rich hexamerins, and the dipteran arylphorins (Beintema et al., 1994). The products encoded by the hexamerin genes are expressed predominantly during late larval life, and thus appear to be negatively regulated by JH. Another group of proteins in the hexamerin superfamily are the hemolymph cyanoproteins (Miura et al., 1998) and the orthopteran hemolymph JH binding proteins (Koopmanschap and deKort, 1988; Braun and Wyatt, 1996). As first demonstrated by Jones et al. (1988, 1990a), the expression patterns of certain fat body genes are downregulated by JH and JH analogs; however, exceedingly high doses of JH analogs were used to elicit the responses. More recently, Hwang et al. (2001) and Cheon et al. (2002) have isolated two genes encoding hexamerins that are JH-suppressible at low doses of JH analog. Expression of these genes is not dose-dependent, but can be downregulated within a 6 h period, suggesting that JH is acting directly on transcription. Thus, a general pattern is emerging that indicates fat body hexamerin expression is suppressed in the presence of JH.

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While few insect endocrinologists would argue with the idea that the fat body is a target for JH, there remain technical difficulties in working with the fat body. Its cellular heterogeneity, massive tracheal penetration, and apparent sensitivity to wounding or to culture in vitro make experimental manipulation of the premetamorphic fat body difficult. More troubling is the fact that many of the studies lack a dose–response curve, and data are based on a single large dose of hormone or analog. As demonstrated by Orth et al. (1999) in their study on JH regulation of hJHBP expression, the effect of increasing doses of JH on the fat body is stimulatory only within a limited range; after peak stimulation is reached, further increases in JH result in a negative response. In this case, a single large dose of hormone would have obscured very important data. Another issue with many of the studies is the lack of authentic controls, especially for experiments in vitro. Large doses of JH can form a monolayer at the surface of the medium, hindering gaseous exchange and compromising the physiology of the fat body in culture. Unless a comparable lipid is used as a control, the results may be meaningless. It is anticipated that technological advances, particularly in molecular analyses, coupled with a more physiological approach to hormone treatment, should ultimately reveal how JH functions with respect to the fat body. 3.7.11.3.4. Muscle responses Myogenesis occurs during two developmental periods: the embryonic period in which larval muscles are developed and the period of larval–pupal metamorphosis, in which adult muscles are formed. In addition to myogenesis, existing larval muscles can undergo cell death, or become restructured and/or reoriented for a new role in the adult (Riddiford, 1994; Roy and Vijay Raghavan, 1999). At the present time there is little evidence to suggest that JH has a direct role on embryonic myogenesis; however, JH involvement in the regulation of muscle development and muscle fate during metamorphosis is well documented (Schwartz, 1992; Riddiford, 1994; Hegstrom et al., 1998; Roy and VijayRaghavan, 1999; Buszczak and Segraves, 2000; Cascone and Schwartz, 2001; Lee et al., 2002). The fate of larval muscles is determined by a finely tuned interplay between JH and ecdysteroids, an example of which is found in the process of proleg retraction in M. sexta (Weeks and Truman, 1986a, 1986b). After the larva wanders in the last larval stadium, the prolegs are no longer required, and muscles that move these structures begin to degenerate. The signal for this process is the commitment peak in the ecdysteroid titer, which occurs in the absence of JH. Under these conditions,

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the retractor muscles become committed to undergo apoptosis (programmed cell death) (see Chapter 2.5), with the actual process of degradation elicited by the large prepupal peak of ecdysteroids that occurs several days later. Despite its concurrent presence at the time of the prepupal peak in the ecdysteroid titer, JH can no longer inhibit apoptosis of the retractor muscles. Adult-specific muscles are formed from specific myoblasts that proliferate and differentiate during the late larval or early pupal stages (Roy and VijayRaghavan, 1999) (see Chapters 2.1 and 2.3). In those muscles that make up the larval ventral diaphragm of M. sexta, proliferation and differentiation are controlled by ecdysteroids, but these two events have different hormonal requirements (Champlin et al., 1999). Moderate levels of ecdysteroids induce proliferation of the ventral diaphragm myoblasts, while high or low doses arrest cellular proliferation in the G2 phase of the cell cycle in these cells. High doses of ecdysteroids, such as those observed at the prepupal peak, induce the myoblasts to exit the cell cycle and express muscle myosin. The ability of critical titers of ecdysteroids to initiate either proliferation or differentiation is modified in the presence of methoprene. For example, high levels of ecdysteroids induce myoblast proliferation, but are unable to induce myoblast differentiation, in the presence of methoprene. These experimental observations correlate well with myoblast development in vivo. The prepupal peak of ecdysteroids, which is sufficient to induce differentiation, is accompanied by a concomitant rise in JH that prevents the myoblasts from differentiating. The myoblasts continue to proliferate until day 7 of the pupal stage when, in the presence of the pupal ecdysteroid peak but in the absence of JH, they then begin to differentiate. While both the prepupal and pupal surges of ecdysteroids are sufficient to cause differentiation, the pupal surge is not accompanied by a rise in JH levels, thus allowing the myoblasts to differentiate. A similar interplay between JH and ecdysteroids is observed in the development of flight muscle in the desert locust, S. gregaria. Flight muscle development is initiated at the beginning of the last nymphal stadium and completed shortly after adult metamorphosis (Wang et al., 1993). In addition to its fully differentiated myofibrils, functional flight muscle contains a high level of a fatty acid binding protein that is involved in sequestering lipids for energy. Flight muscle development begins during the fifth nymphal stadium, when JH is absent and ecdysteroid titers are high. When a large dose of methoprene is applied to the last-stadium nymph, the insect

undergoes another nymphal molt, and flight muscles that should have formed working myofibrils remain undifferentiated and lack fatty acid binding protein. Thus, elevated JH titers in the last nymphal stadium can lead to either retention of nymphal muscle or delay in the development of adult muscles. In the adult stage, JH can have the opposite effect on muscles, inducing them to undergo degradation to yield precursors for vitellogenin production (Rose et al., 2001). Treating the alate or winged form of the aphid Acyrthosiphon pisum with a JH analog initiates histolysis of indirect flight muscles via the ubiquitin-dependent pathway for apoptosis (Kobayashi and Ishikawa, 1994). Conversely, treatment with precocene II, a compound that reduces JH biosynthesis in this species, prevents flight muscle breakdown. A similar situation is found in another aphid species, A. fabae (Hardie et al., 1990). The development of flight muscle in Orthoptera appears to follow this same pattern, i.e., JH either induces and/or is involved in the histolysis of these muscles. Application of JH III or methoprene to the cricket Modicogryllus confirmatus induces the degeneration of flight muscles within 3 days (Tanaka, 1994). In addition to JH, there may be neural factors that are required for the degeneration of specific flight muscles (Shiga et al., 2002). 3.7.11.3.5. Prothoracic gland responses It has long been known that the prothoracic glands, the site of synthesis of ecdysteroids, are regulated by ecdysone itself, via short positive and negative feedback loops (Williams, 1952; Siew and Gilbert, 1971; Sakurai and Williams, 1989) (see Chapter 3.8). Given the close relationship between JH and ecdysteroids in regulation of metamorphosis, the effect of JH on the prothoracic glands has also been examined (Gilbert, 1962; Siew and Gilbert, 1971). JH has two separate and distinct effects: prevention of precocious degeneration of the glands and regulation of ecdysteroid synthesis/secretion during the larval stage. In M. sexta, the prothoracic glands undergo apoptosis between 5 and 6 days after pupation, at the time of the pupal peak in the ecdysteroid titer. Dai and Gilbert (1999) demonstrated that early pupal glands, in the presence of ecdysteroids in vitro, will also undergo apoptosis. Glandular degeneration can be delayed for up to a week, if a newly molted pupa is injected with a large dose of JH II. Moreover, the glands remain capable of synthesizing ecdysone (Dai and Gilbert, 1998). The underlying mechanisms by which JH prevents this important developmental event is still unclear. Nijhout (1994), in his elegant review of JH effects on ecdysteroid synthesis, came to the conclusion

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that JH does not act directly on the prothoracic glands to modulate ecdysone biosynthesis. His model suggests that elevated levels of JH during the early part of the last larval stadium act on the brain (or other intermediary tissue) to suppress synthesis or release of PTTH, the hormone responsible for stimulating ecdysteroidogenesis in the prothoracic glands. It is only when the JH titers drop that PTTH can be released to initiate commitment to pupation by stimulating synthesis of the commitment peak in the ecdysteroid titer (Nijhout and Williams, 1974; Rountree and Bollenbacher, 1986). The idea that the brain is the major target is supported by the work of Gruetzmacher et al. (1984), who demonstrated that although PTTH could stimulate M. sexta prothoracic gland activity in vitro, none of the JH homologs or methoprene, even at a number of different doses, were able to do so. Evidence presented by Sakurai et al. (1989) suggests that JH not only acts on the brain but also on the glands themselves, to directly suppress the acquisition of competence to respond to PTTH. Thus, under normal physiological conditions in the last larval stadium, the JH-regulated inhibition of PTTH secretion and suppression of gland competence probably evolved as a safety mechanism to prevent the premature onset of the metamorphic molt (Nijhout, 1994). While it is accepted that high levels of JH inhibit PTTH production or secretion during the feeding period of the last larval stadium, this does not appear to be the case either in the earlier stadia or after commitment to pupation. In the earlier stadia, JH titers are high to maintain the larval state. Presumably, these high levels of JH should inhibit PTTH secretion, yet the insects undergo normal molts. In early studies, the implantation of extra CA into penultimate and last instars of G. mellonella invariably led to perfect supernumerary molts, often more than one (Granger and Sehnal, 1974; Sehnal and Granger, 1975). In more recent studies, supplementing normal JH titers with an analog, fenoxycarb, did not block third and fourth instars of B. mori from undergoing normal molts to fourth and fifth instars; however, an extra molt to a perfect sixth instar was noted (Kamimura and Kiuchi, 2002). An identical result was seen in M. sexta using methoprene (Lonard et al., 1996). An in vitro study provides a basis for the results of these in vivo experiments. Addition of fenoxycarb to incubations in vitro of B. mori prothoracic glands taken throughout the last stadium results in an extreme suppression of ecdysteroidogenesis (Dedos and Fugo, 1996). Although a recent report has shown that JH I, applied in large doses early in the last

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larval stadium, can transiently elevate PTTH levels in the hemolymph of B. mori (Mizoguchi, 2001), the majority of the studies done so far indicate that in contrast to the model for JH–ecdysteroid interaction during larval–pupal metamorphosis, JH is capable of acting directly on the prothoracic glands and does so by suppressing ecdysteroid synthesis. Biochemical evidence supports this contention: hydroprene, a JH analog, and JH I significantly delay the ability of the prothoracic glands to respond to PTTH or MIX, an inhibitor of cyclic nucleotide phosphodiesterase (Gu et al., 1997). Since PTTH activates an intracellular second messenger system involving cyclic nucleotides (Gilbert et al., 2000) (see Chapter 3.2), MIX can serve as a surrogate for PTTH by inhibiting their catabolism. The fact that neither PTTH nor MIX can activate ecdysteroidogenesis in the presence of JH strongly reinforces the idea that JH can act directly on the glands. Several groups have demonstrated that the effect of JH on the prothoracic glands changes after commitment to pupation. Insects that have been debrained or neck-ligated to remove the source of PTTH respond to JH or analog treatment with increased ecdysteroid synthesis (Hiruma et al., 1978; Safranek et al., 1980; Gruetzmacher et al., 1984; Dedos and Fugo, 1996; Dai and Gilbert, 1998). Thus, a reversal in hormone action appears to have taken place at, and possibly as a result of, commitment. As we have seen with other tissues, the response to JH can vary depending upon the developmental period. In the case of the prothoracic glands, the regulatory processes that control ecdysteroidogenesis and involve JH are very complex.

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3.7.12. Molecular Mode of Action of the Juvenile Hormones No sooner had the chemical structure of JH I been reported by Ro¨ ller et al. (1967), one of the first of many reviews on JH action at the molecular level appeared in print (Kroeger, 1968). Since that time, there have been a number of major reviews focusing on the topic, with at least nine appearing during the last dozen years (Kumaran, 1990; Riddiford, 1994, 1996; Jones, 1995; Wyatt and Davey, 1996; Gade et al., 1997; Gilbert et al., 2000; Lafont, 2000; Wheeler and Nijhout, 2003). Despite the volumes written about the subject, the molecular action of JH remains very much a mystery. This has led some to suggest that JH may have multiple roles in cellular and molecular mechanisms (Jones, 1995; Wheeler and Nijhout, 2003). Our assessment of the literature suggests that some of the problems in deciphering the molecular action

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of JH lie not in unfathomable molecular mechanisms, but rather in the quality of the hormone, choice of experimental model, and development of experimental design. A major experimental pitfall is the quality of the hormone, especially commercially available JH III from a US supplier. The supplier acknowledges that the hormone is racemic and the level of contamination is significant. Yet there appears to be a tacit assumption among investigators that the unnatural racemate and contaminants have no effect on the experimental procedure, an assumption that is unwarranted until proven. Long-term storage of the hormone and its metabolites may also be problematic and a means of repurifying the compounds must be available. A similar criticism can be leveled at the use of commercial-grade JH agonists that may contain significant amounts of incipients with unknown biological functions. Metamorphosis is undoubtedly one of the most striking phenomena in the biological world, yet it has not yielded its molecular secrets easily, particularly with regard to JH action. While major strides have been made in the last few years, using a paradigm that depends on the absence of JH makes dissecting the molecular action of the hormone challenging. This problem is further compounded by the fact that the metamorphic actions of JH on cellular and molecular events often require lengthy incubation periods. These long periods make separating key regulatory events from secondary events difficult. It may well be that the extreme complexity of metamorphosis does not lend itself well to isolating the actions of JH, and, as Jones (1995) and Wheeler and Nijhout (2003) note, the hormone may act on a number of molecular pathways in the same cell. As our knowledge of larval physiology and molecular biology expands, nonmetamorphic events regulated by JH, such as pigmentation and metabolic processes, may provide more tractable experimental models with which to study the molecular actions of JH at the genomic level. s0480

3.7.12.1. Juvenile Hormone Interaction with Ultraspiracle

One of the more intriguing discoveries since the last major reviews is that presented by Jones and Sharp (1997), who demonstrated that the nuclear receptor Ultraspiracle (USP) can bind JH III (see Chapter 3.5). USP, an orphan receptor for which no endogenous ligand has been unambigiously established (Billas et al., 2001), is a heterodimeric partner that interacts with ECR to form the functional ECR complex (Thomas et al., 1993; Yao et al., 1993) or

with DHR38, the D. melanogaster ortholog of the mammalian NGFI-B subfamily of orphan nuclear receptors (Baker et al., 2003) (see Chapter 3.5). USP is an ortholog of the vertebrate retinoid X receptor (RXR) (Oro et al., 1990), which is responsive to very high levels of methoprene (Harmon et al., 1995). Using a fluorescence assay that detects changes in protein conformation induced by ligand binding, Jones and Sharp (1997) demonstrated that JH III and JH III acid change the conformation of recombinant D. melanogaster USP, while farnesol and 20E do not. The USP–JH III acid interaction generates anomalous spectra, suggesting that it may not trigger a conformational change that is identical to that obtained with JH III. These investigators estimate the dissociation constant of the USP: JH III complex to be approximately 1 mM, a value considerably higher than might be expected for a nuclear receptor that must sequester hormone directly from the circulatory system. Further refinement of the original studies indicated that methoprene competes with JH III for a binding site on USP (Jones et al., 2001). It should be cautioned that these studies used only partially purified (75%) racemic JH III, yet the investigators chose to express ligand concentrations as ‘‘concentrations of the natural active isomer in the respective binding reaction’’ (Jones et al., 2001). Arbitrarily overlooking the impurities in the hormone preparation and using an estimated 10R JH III concentration may present problems in the interpretation of the binding constants. Nevertheless, evidence from functional transcription assays supports this model. Using modified USP response elements coupled to a JHE core promoter (61 to þ28 relative to the start site) and a reporter gene, Jones et al. (2001) demonstrated that construct expression could be induced by JH III in a dose-dependent fashion. The promoter construct was only weakly responsive to all trans-retinoic acid and did not respond to triiodo-l-thyronine (Jones et al., 2001). Functional transcriptional assays also demonstrate that point mutations in the putative JH III binding site of USP, which abolish JH binding, cause the mutant receptor to act as a dominant negative and suppress JH III activation (Xu et al., 2002). Jones et al. (2001) suggested that high levels of JH III induce a conformational change in structure to stabilize dimeric/oligomeric forms of USP. This interaction may be important in dimerization of USP with its ecdysteroid receptor partner, but how it occurs and the role of USP–DNA binding in stabilization of the complex is not yet understood. It is well known that ecdysteroids induce enhanced

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heterodimer formation even when the binding of the heterodimers to cognate DNA response elements does not occur (Lezzi et al., 2002). The initial crystallographic analyses of the USP ligand binding domains from H. virescens (Billas et al., 2001) and D. melanogaster (Clayton et al., 2001) indicated that the putative ligand binding site is locked in an antagonist conformation. More recently, Sasorith et al. (2002) reinvestigated the site and found that recombinant USP could bind ligands and attempted, using computer algorithms, to dock putative ligands in the binding site. Their studies indicate that the ligand binding site is relatively large and predominantly lined with hydrophobic residues. Interestingly, the binding site of the recombinant USP contains a phospholipid inserted by the expression host, and it would be of considerable interest to know how this lipid can affect JH binding. Computation docking of JH and analogs in the binding site of recombinant H. virescens USP suggests that JH may fit the putative site; however, the percentage of occupancy of the ligand binding sites by these ligands lies in the bottom range of values for classical nuclear receptors, thus raising concern about the validity of USP as the juvenile hormone receptor (Sasorith et al., 2002). USP, with its large ligand binding site and a low level of occupancy, behaves more like a sensor than a classical high affinity receptor. Indeed, using the sensor model (Chawla et al., 2001), one might envision an entirely different role for USP, based on nutritional levels in the premetamorphic larva. When sufficient levels of a key dietary lipid(s) have been attained, the USP binding site becomes saturated with that nutritional ligand, which, in turn, alters the conformation of the protein. Ligand-activated USP might then enhance ecdysteroid binding by its partner, ECR. Ligand activation of the appropriate ecdysteroid receptor isoforms leads to changes in neuronal activity (Hewes and Truman, 1994; Truman, 1996) that may trigger activity in the prothoracicotropes or dendritic fields enervating them. While this hypothesis is simplistic, it could account for the very early signaling events that initiate ecdysis. It now seems clear that USP binds JH, albeit with low affinity, as well as other lipids. It may be that refinement of experiments presented by the Jones laboratory will shed new light on how the hormone acts at the molecular level. 3.7.12.2. Juvenile Hormone Interaction with the Methoprene-Tolerant Gene

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Using mutagenesis studies to dissect the molecular action of JH, Wilson and his group have examined a

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locus in D. melanogaster, the Methoprene-tolerant (Met) locus, that encodes for a protein similar in sequence to proteins in the basic-helix–loop–helix– PAS (bHLH-PAS) family of transcriptional regulators (Wilson and Ashok, 1998). Treating wild-type flies with high levels of methoprene at certain times during larval development leads to a distinct array of developmental abnormalities, including aberrations in the central nervous system, salivary glands, and muscles (Restifo and Wilson, 1998). In contrast to wild-types, flies carrying the Met mutation are resistant to high levels of methoprene and appear to develop and reproduce normally (Wilson and Fabian, 1986). The Met gene (GenBank Accession no. AF034859) encodes for a 716-residue protein which in wild-type flies displays a dissociation constant of 6 nM for JH III (Shemshedini et al., 1990). Competitive displacement studies comparing the homologs indicate that JH I, JH II, JH III acid, and methoprene are weak competitors when compared with JH III; methoprene binding is 100-fold weaker than that of JH III. Flies carrying the Met mutation exhibited a significantly reduced (sixfold) binding affinity for JH III (38 nM). The reduced affinity of the 85 kDa Met protein for methoprene translates into a 50–100-fold increase in resistance to both its toxic and morphogenetic effects. Flies carrying the Met mutation are also resistant to toxic levels of the naturally occurring hormones JH III and JHB3, but not to various other classes of insecticides. This important distinction demonstrates that Met is not a general insecticide-resistance gene, but is specific for JH and JH analogs (Wilson et al., 2003). Analysis of the Met sequence indicates it has three regions displaying similarity to the bHLH–PAS family of transcriptional regulators. It is instructive to note that the bHLH–PAS gene family was named for three important members of the group: the Period gene, the Aryl hydrocarbon receptor gene, and the Single-minded gene. Of particular importance is the similarity of Met to the Aryl hydrocarbon receptor (Ahr) gene that encodes for a xenobiotic binding protein (Ashok et al., 1998). The fact that methoprene interacts with a member of the AHR family of receptors may explain why so many compounds structurally distant from JH have JH-like activity (Stahl, 1975). With its similarity to AHRs, Met could bind hydrophobic ligands that marginally resemble JH and regulate certain JH-responsive genes (Ashok et al., 1998). Equally interesting is the fact that the vertebrate AHR partners with another protein, the aryl hydrocarbon nuclear translocator, to form an active transcriptional regulator that activates genes responsible for mediating the

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response to xenobiotics. While no partner has yet been found for Met, the potential for such an interaction has been noted (Restifo and Wilson, 1998). Immunolocalization studies suggest that Met is limited to nuclei (Pursley et al., 2000), thus ruling out the earlier suggestion that Met acts as a cytosolic ‘‘sink’’ (Shemshedini et al., 1990). Developmentally, it is present in all cells of the Drosophila embryo from the 256 cell stage until early gastrulation, when the signal begins to decline. In second and third stadium larvae Met is found in salivary glands, fat body, imaginal discs, and gut primordium. Met is also present in pupal histoblasts and in adult reproductive tissues, including the female ovarian follicle cells and spermatheca and the male accessory glands (Pursley et al., 2000). To date, over a dozen alleles of Met have been recovered in genetic screens (Wilson et al., 2003), but one in particular, Met27, poses a perplexing question about the role of Met in JH action. When examined by Northern analysis and RT-PCR, flies homozygous for Met27 lack Met transcripts, yet these insects survive and develop into adults. The lack of Met expression becomes evident only in the adult female, in which there is an 80% reduction in oogenesis (Wilson and Ashok, 1998). If flies carrying a null mutation in Met can undergo seemingly normal development and limited oogenesis, both events being under JH control, the role of Met in JH action is open to question. Pursley et al. (2000) suggest that ‘‘genetic redundancy’’ (Krakauer and Plotkin, 2002), the equivalent of back-up systems for biological phenomena, may be involved in this situation. Given the apparent lack of Met functions during the larval stages, but its significant role in both male and female reproduction (Wilson and Ashok, 1998; Wilson et al., 2003), it may be that JH utilizes several different and distinct molecular pathways to control development and reproduction. 3.7.12.3. Juvenile Hormone and Metamorphosis: The Role of the Broad-Complex

A growing body of evidence has implicated the transcription factor Broad (FlyBase identification no. FBgn0010011) as a key component in the initiation of metamorphosis (Riddiford et al., 2003). Mutations in Broad, a member of the Broad-Tramtrack-Bric-a-Brac family, allow normal larval development to occur but prevent pupation (Kiss et al., 1988; Bayer et al., 1996; Riddiford et al., 2003). Less severe Broad phenotypes have wings that are shorter and wider than wild-types. The Broad gene is large (100 kb) and in Drosophila encodes for at least six isoforms of the protein,

which all share a common N-terminal of approximately 425 amino acids (Bayer et al., 1996). Through alternative splicing, the C-terminal can be represented by any one of four pairs of C2H2-type zinc finger domains, Z1, Z2, Z3, and Z4. The Nand C-terminals are connected by linkage domains of varying lengths. The Z4 transcript encodes a protein of 877 amino acids. After the onset of metamorphosis, the nuclei of all larval and imaginal cells display Broad; however, each tissue appears to have its own specific constellation of isoforms that appear in a temporally unique sequence (Mugat et al., 2000). For example, Mugat et al. (2000), using an isoform-specific antibody in their studies on D. melanogaster fat body, followed a shift from Z2 to the other isoforms as metamorphic events proceed. Bayer et al. (1996) demonstrated that Z1 is the predominant isoform during pupal cuticle formation in the abdominal epidermis and imaginal discs. Ingestion of the JH analog pyriproxifen by first instar D. melanogaster prolongs the third stadium by several days (Riddiford et al., 2003) and results in a 12 h delay in the appearance of Broad in abdominal epidermis. By the time of pupation, however, Broad protein levels were similar to controls, indicating that this potent JH analog cannot prevent the appearance of this protein. The adult abdominal epidermis, formed from nests of tissue called histoblasts, begins to proliferate and spread over the abdomen after puparium formation. These new cells displace the existing larval cells. Broad is found in larval cells that are destined to die, but cannot be found in imaginal cells once they have spread across the abdomen. Interestingly, JH treatment delays the loss of Broad in the imaginal cells and causes these cells to produce proteins indicative of a pupal-like cuticle. Increasing Z1 levels through misexpression at the onset of adult cuticle formation mimics the effects of JH, causing the reappearance of mRNAs encoding pupal cuticular proteins and the suppression of mRNAs encoding adult cuticular proteins (Zhou and Riddiford, 2002). These investigators suggest that JH prolongs the expression of the Z1 isoform, and suppresses the onset of adult cuticular formation. In M. sexta, only three Broad isoforms have been discovered, Z2, Z3, and Z4, with Z4 being most prominent in epidermal tissue involved in pupal cuticle formation (Zhou et al., 1998; Zhou and Liu 2001). As observed in D. melanogaster, the protein’s appearance is well-defined spatially and temporally, following the pattern of pupal commitment, as defined by the loss of sensitivity to JH. Most significantly, within 6 h of 20E treatment, epidermal

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preparations in vitro display a marked rise in Broad mRNA (Zhou et al., 1998; Zhou and Riddiford, 2002). This rise signals the onset of commitment to pupation. In cells that are not yet committed, JH I prevents the induction of Broad mRNA synthesis by 20E (Zhou et al., 1998). Once Broad appears in the cells, JH can no longer suppress the accumulation of Broad mRNA, and its levels remain elevated, even during the rise in JH titers later in the stadium. Broad expression eventually disappears during the early pupal stage coinciding with the onset of adult cuticular synthesis. The role of Broad was further defined by applying sufficient JH to pupae to elicit the formation of a second pupal cuticle; in this case, Broad mRNA and protein rose after exposure to 20E, then dropped after the second pupal cuticle was formed (Zhou and Riddiford, 2002). Thus, Broad’s presence in the cell appears to inhibit larval cuticular synthesis, while permitting pupal cuticular synthesis to proceed. Moreover, Broad appears to suppress the synthesis of the adult cuticle. Exogenous JH can delay but not prevent the expression of Broad, but the molecular actions of the hormone remain unclear. 3.7.12.4. Juvenile Hormone Regulation of Cytosolic Malate Dehydrogenase p1320

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JH has long been implicated in the regulation of cellular energy production and of biochemical and second messenger pathways. This long history includes research on such diverse mechanisms as modification of oxidative phosphorylation (Clarke and Baldwin, 1960; Chefurka, 1978); shifts in cellular potassium levels (Kroeger, 1968); membrane perturbation (Baumann, 1969; Barber et al., 1981); second messenger regulation (Everson and Feir, 1976; Kensler et al., 1978; Yamamoto et al., 1988); changes in cytoskeletal structure (Capella and Hartfelder, 2002); and the development of asymmetric organs (Adam et al., 2003). Interest in certain of these areas has been rekindled by a series of studies carried out by Farkas and his associates on the regulation of cytosolic malate dehydrogenase in D. melanogaster (Farkas and Knopp, 1997, 1998; Farkas and Sutakova, 2001; Farkas et al., 2002). Cytosolic malate dehydrogenase (EC 1.1.1.40), also known as NADP-malic enzyme (ME), is involved in production of cytoplasmic NADPH. The enzyme is particularly prevalent in tissues that are involved in fatty acid synthesis and it is a target for various hormones in vertebrates (Farkas et al., 2002). In a series of papers, Farkas and his group have demonstrated that ME, a product of the Men gene (CG 10120; FBgn 0002719), is under the

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regulation of JH during the mid-third stadium of D. melanogaster. A dose of 5 pg of JH III will elicit half-maximal ME activity and while this level is higher than physiological levels (0.02 pg per larva) (Sliter et al., 1987), it is still lower than the dose typically used to induce morphological abberations (Riddiford and Ashburner, 1991). The response to JH III occurs in two phases, an early phase occurring 3 h after treatment, in which ME activity doubles, and a late phase occurring 12 h after treatment, in which a threefold increase over the control is observed. The transcriptional inhibitor actinomycin D does not block the early ME rise, but does block the late rise, suggesting that JH can increase ME activity without new protein synthesis. That JH added directly to the cytosol can not induce the ME rise indicates that the hormone may be acting through a pre-existing pathway that is immediately affected, such as that of a second messenger cascade (Farkas et al., 2002). In contrast to the early rise, the late rise in ME activity is sensitive to actinomycin D, indicating that this rise is dependent on de novo transcription of ME. The authors have yet to examine the levels of ME mRNA that would confirm the inhibitor studies. Using developmentally staged, wild-type larvae and the temperature-sensitive mutants ecd1 and su(f)ts67g, Farkas et al. (2002) demonstrated that ecdysteroids are also involved in the JH-regulated ME rise. Wild-type flies respond to JH only near the time of pupariation, when ecdysteroid titers begin to rise. By contrast, in temperature-sensitive, ecdysone-deficient mutants, JH is unable to initiate the expected ME rise at elevated temperatures. These observations suggest that a complex synergistic regulatory mechanism is acting to control ME levels. A low level of ecdysteroid must be present before JH can induce activation of ME, yet high levels of ecdysteroid, such as those seen at pupariation, downregulate the activity of ME. The physiological role of JH in upregulating this event cannot be well understood unless placed in the context of lipogenesis. Farkas et al. (2002) suggest that the ME–JH interaction may play a role in increasing the levels of unsaturated fatty acids. JH and certain JH analogs have been shown to induce a rise in unsaturated fatty acids (Schneider et al., 1995) and phospholipid levels (Della-Cioppa and Engelmann, 1984) in adult S. gregaria and L. maderae. In D. melanogaster, the ecdysteroid-induced onset of metamorphosis leads to the cessation of feeding; it may be that high levels of JH during the earlier stadia and the early part of the last stadia ensure availability of lipid resources during metamorphosis. While the pathways are still not clear, this paradigm of JH

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action at the molecular level holds much promise for better understanding how the hormone may regulate homeostatic mechanisms that are necessary for insect growth and development.

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Mas-AS immunoreactive peptide in Lacanobia oleracea. Insect Biochem. Mol. Biol. 28, 775–784. Audsley, N., Weaver, R.J., Edwards, J.P., 1999a. Juvenile hormone synthesis by corpora allata of tomato moth, Lacanobia oleracea, and the effects of allatostatins and allatotropin in vitro. Eur. J. Entomol. 96, 287–297. Audsley, N., Weaver, R.J., Edwards, J.P., 1999b. Significance of Manduca sexta allatostatin in the tomato moth Lacanobia oleracea. In: Sandman, C.A., Strand, F.L., Beckman, B., Cronwall, B.M., Flynbn, F.W., Nachman, F.W. (Eds.), Neuropeptides, Structure and Function in Biology and Behavior, vol. 897. New York Academy of Science, New York, pp. 330–341. Audsley, N., Weaver, R.J., Edwards, J.P., 2000. Juvenile hormone biosynthesis by corpora allata of larval tomato moth, Lacanobia oleracea, and regulation by Manduca sexta allatostatin and allatotropin. Insect Biochem. Mol. Biol. 30, 681–689. Audsley, N., Weaver, R.J., Edwards, J.P., 2002. Degradation of Manduca sexta allatostatin and allatotropin by proteases associated with the foregut of Lacanobia oleracea larvae. Peptides 23, 2015–2023. Auerswald, L., Birgul, N., Gade, G., Kreienkamp, H.J., Richter, D., 2001. Structural, functional, and evolutionary characterization of novel members of the allatostatin receptor family from insects. Biochem. Biophys. Res. Commun. 282, 904–909. Baehr, J.-C., Caruelle, J.P., Poras, M., 1986. The activity of denervated corpora allata in a diapausing strain of Locusta migratoria: in vivo and in vitro studies. Int. J. Invertebr. Reprod. Devel. 10, 143–150. Baehr, J.-C., Porcheron, P., Papillon, M., Dray, F., 1979. Haemolymph levels of juvenile hormone, ecdysteroids and protein during the last two larval instars of Locust migratoria. J. Insect Physiol. 25, 371–460. Baehr, J.-C., Pradelles, P., Lebreux, C., Cassier, P., Dray, F., 1976. A simple and sensitive radioimmunoassay of insect juvenile hormone using an iodinated tracer. FEBS Lett. 69, 123–128. Baker, F.C., 1990. Techniques for identification and quantification of juvenile hormones and related compounds in arthropods. In: Gupta, A.P. (Ed.), Morphogenetic Hormones of Arthropods. Rutgers University Press, New Brunswick, NJ, pp. 389–444. Baker, F.C., Lanzrein, B., Miller, C.A., Tsai, L.W., Jamieson, G.C., et al., 1984. Detection of only JH III in several life-stages of Nauphoeta cinerea and Thermobia domestica. Life Sci. 35, 1553–1560. Baker, K.D., Shewchuk, L.M., Kozlova, T., Makishima, M., Hassell, A., et al., 2003. The Drosophila orphan nuclear receptor DHR38 mediates an atypical ecdysteroid signaling pathway. Cell 113, 731–742. Baker, F.C., Tsai, L.W., Reuter, C.C., Schooley, D.A., 1987. In vivo fluctuation of JH, JH acid, and ecdysteroid titer, and JH esterase activity, during development of fifth stadium Manduca sexta. Insect Biochem. 17, 989–996. Baker, F.C., Tsai, L.W., Reuter, C.C., Schooley, D.A., 1988. The absence of significant levels of the known

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Woodring, J., Hoffmann, K.H., 1994. The effects of octopamine, dopamine and serotonin on juvenile hormone synthesis, in vitro, in the cricket, Gryllus bimaculatus. J. Insect Physiol. 40, 797–802. Wroblewski, V.J., Harshman, L.G., Hanzlik, T.N., Hammock, B.D., 1990. Regulation of juvenile hormone esterase gene expression in the tobacco budworm. Arch. Biochem. Biophys. 278, 461–466. Wyatt, G.R., Davey, K.G., 1996. Cellular and molecular actions of juvenile hormone. 2. Roles of juvenile hormone in adult insects. Adv. Insect Physiol. 26, 1–155. Xu, Y., Fang, F., Chu, Y.X., Jones, D., Jones, G., 2002. Activation of transcription through the ligand-binding pocket of the orphan nuclear receptor ultraspiracle. Eur. J. Biochem. 269, 6026–6036. Yagi, K.J., Tobe, S.S., 2001. The radiochemical assay for juvenile hormone biosynthesis in insects: problems and solutions. J. Insect Physiol. 47, 1227–1234. Yagi, K.J., Konz, K.G., Stay, B., Tobe, S.S., 1991. Production and utilization of farnesoic acid in the juvenile hormone biosynthetic pathway by corpora allata of larval Diploptera punctata. Gen. Comp. Endocrinol. 81, 284–294. Yagi, S., Kuramochi, K., 1976. The role of juvenile hormone in larval duration and spermiogenesis in relation to phase variation in the tobacco cutworm, Spodoptera litura. Applic. Entomol. Zool. 11, 133–138. Yamamoto, C.K., Chadarevian, A., Pellegrini, M., 1988. Juvenile hormone action mediated in male accessory glands of Drosophila by calcium and kinase C. Science 239, 916–919. Yang, J.H., Park, J.-C., Lee, B.H., 2000. Molecular characterization of allatostatins cDNA in central nervous system and midgut from the German cockroach Blattella germanica. Korean J. Entomol. 30, 131–137. Yao, T.P., Forman, B.M., Jiang, Z.Y., Cherbas, L., Chen, J.D., et al., 1993. Functional ecdysone receptor is the product of EcR and ultraspiracle genes. Nature 366, 476–479. Yin, C.-M., Zou, B.X., Jiang, M.G., Li, M.F., Qin, W.H., et al., 1995. Identification of juvenile hormone III bisepoxide (JHB3), JH III and methyl farnesoate secreted by the corpus allatum of Phormia regina, in vitro and function of JHB3 either applied alone or as a part of a juvenoid blend. J. Insect Physiol. 41, 473–488.

Yin, C.-M., Takeda, M., Wang, Z.-S., 1987. Juvenile hormone analogue, methoprene as a circadian and development modulator in Diatraea grandiosella. J. Insect Physiol. 33, 95–102. Young, J.K., Orth, A.P., Goodman, W.G., 2003. Allelic variation in the hemolymph juvenile hormone binding protein gene of Manduca sexta. Mol. Cell. Endocrinol. 208, 41–50. Yu, C.G., Hayes, T.K., Strey, A., Bendena, W.G., Tobe, S.S., 1995. Identification and partial characterization of receptors for allatostatins in brain and corpora allata of the cockroach Diploptera punctata using a binding assay and photoaffinity labeling. Regul. Peptides 57, 347–358. Yu, S.J., Terriere, L.C., 1978. Metabolism of juvenile hormone I by microsomal oxidase, esterase and epoxide hydratase of Musca domestica and some comparisons with Phormia regina and Sarcophaga bullata. Pestic. Biochem. Physiol. 9, 237–246. Zera, A.J., Sanger, T., Hanes, J., Harshman, L., 2002. Purification and characterization of hemolymph juvenile hormone esterase from the cricket, Gryllus assimilis. Arch. Insect Biochem. Physiol. 49, 41–55. Zhou, B.H., Hiruma, K., Shinoda, T., Riddiford, L.M., 1998. Juvenile hormone prevents ecdysteroid-induced expression of broad complex RNAs in the epidermis of the tobacco hornworm, Manduca sexta. Devel. Biol. 203, 233–244. Zhou, C., Liu, B., 2001. Identification and characterization of a silk gland-related matrix association region in Bombyx mori. Gene 277, 139–144. Zhou, X.F., Riddiford, L.M., 2002. Broad specifies pupal development and mediates the ‘‘status quo’’ action of juvenile hormone on the pupal–adult transformation in Drosophila and Manduca. Development 129, 2259–2269. Ziese, S., Dorn, A., 2003. Embryonic integument and ‘‘molts’’ in Manduca sexta. J. Morphol. 255, 146–161. Zimowska, G., Rembold, H., Bayer, G., 1989. Juvenile hormone identification, titer, and degradation during the last larval stadium of Spodoptera littoralis. Arch. Insect Biochem. Physiol. 12, 1–14. Zˇ itnˇ an, D., Kingan, T.G., Kramer, S.J., Beckage, N.E., 1995. Accumulation of neuropeptides in the cerebral neurosecretory system of Manduca sexta larvae parasitized by the braconid wasp Cotesia congregata. J. Comp. Neurol. 356, 83–100.

3.8 Feedback Regulation of Prothoracic Gland Activity S Sakurai, Kanazawa University, Kanazawa, Japan ß 2005, Elsevier BV. All Rights Reserved.

3.8.1. Introduction 3.8.2. Feedback Regulation of Brain–Prothoracic Gland Axis 3.8.2.1. Feedback Regulation of Brain Neurosecretory Cells by 20-Hydroxyecdysone 3.8.2.2. Feedback Regulation of PTTH Cells by 20-Hydroxyecdysone 3.8.3. Feedback Regulation of the Prothoracic Gland by Ecdysteroid 3.8.3.1. Feedback Activation of Ecdysteroid Production 3.8.3.2. Feedback Inhibition 3.8.3.3. Complete and Incomplete Inactivation 3.8.4. Mechanisms of Decrease in Hemolymph Ecdysteroid Titer: Biochemical Aspects 3.8.4.1. Inactivation of 20-Hydroxyecdysone 3.8.4.2. Suppression of Ecdysteroidogenesis 3.8.5. Molecular Mechanism of Feedback Inhibition 3.8.5.1. Feedback Mediated by the EcR/USP Heterodimer 3.8.5.2. Acute and Delayed Inhibition 3.8.6. Horizontal Control of the Prothoracic Gland 3.8.6.1. Effects of JH on PTTH Secretion 3.8.6.2. Effects of JH on Ecdysteroidogenesis 3.8.6.3. Growth-Blocking Peptide–Corpora Allata–Prothoracic Gland Pathway 3.8.7. Role of Feedback Regulation in Larval Growth and Metamorphosis: Meaning of the Shapes of the Ecdysteroid Peak 3.8.8. Future Directions

3.8.1. Introduction p0005

The physiologic effects of insect hormones depend to a large extent on their concentration in the hemolymph. The hormone concentration exposed to target cells is determined by three factors: synthesis and secretion, the downregulation of hormone secretion, and the rates of degradation and elimination. The synthesis, secretion, and downregulation are to a great extent mediated by positive and negative feedback circuits. In the vertical sequences like the hypothalamic–pituitary–thyroid axis of adult vertebrates for example, thyroid hormone suppresses the synthesis and release of thyrotropin releasing hormone (TRH) from neurosecretory cells (NSCs) in the paraventricular nucleus of the hypothalamus as well as thyroid stimulating hormone (TSH) from the thyrotrophs in the adenohypophysis. The control of hypothalamic TRH release by thyroid hormone is referred to as long-loop feedback, and that of TSH release from the pituitary gland as short-loop feedback, while the control of the thyroid gland itself by thyroid hormone is called ultra-short-loop feedback. The definition of feedback is, however, rather vague among insect endocrinologists and

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the term ‘‘feedback’’ has been used occasionally to denote a ‘‘horizontal’’ connection between two or more secretory organs, i.e., the reciprocal interaction between the prothoracic gland and corpus allatum, although these organs are not organized vertically. In the present chapter, the term feedback indicates both feedback in the vertical sequences and horizontal connections, thus covering an intricate network. In addition, the terms, long-loop, short-loop, and ultra-short-loop will not be used, those being simply denoted as feedback unless otherwise indicated. The endocrine control in insects governs diverse biological events in gamete formation, embryonic development, postembryonic development, homeostasis, and behavior, many of which are believed to be under the control, directly or indirectly, of neuropeptides, and therefore neurosecretion. The insect endocrine system is significantly neuroendocrine, as nearly 17% of the cells of the central nervous system (CNS) produce neuropeptides (Ga¨de et al., 1997; Predel and Eckert, 2000) (see Chapter 3.10). Although each ganglion in the CNS contains a number of NSCs with different characteristics with regard to

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p0015

both size and developmental changes in their shapes and content, only a few NSCs have been characterized on the basis of their product. Even in the secretory cells whose products have been identified and their roles established, very little is known about the feedback regulation of cell activity. In insects, a few endocrine glands are part of the peripheral secretory organ system. These are the prothoracic gland that secretes ecdysone (see Chapter 3.3), the corpus allatum that secretes juvenile hormone (JH) and functions as a neurohemal organ for various neuropeptides produced by the brain NSCs (see Chapter 3.7), the corpus cardiacum that secretes peptide hormones involved in the control of energy metabolism (Ga¨de et al., 1997), the epitracheal gland that includes the Inka cell that releases ecdysis triggering hormone (Zitnan et al., 1996) (see Chapter 3.1), the ovary and testes, both of which are known to produce ecdysone in a distinct period of the insect life cycle (Hagedorn et al., 1975; Lagueux et al., 1977; Hetru et al., 1978; Delbecque et al., 1990) (see Chapter 3.9), the gut that contains a number of myotropic factors (Ga¨ de et al., 1997) (see Chapter 4.5), epidermal cells, a source of dopamine (Noguchi and Hayakawa, 1996) and ecdysone in the argasid tick (Zhu et al., 1991) (see Chapter 3.16), and the fat body secreting an insect cytokine, growth-blocking peptide (Hayakawa et al., 1998) and growth factors, one of which is imaginal disc growth factor (IDGF) known to stimulate cell proliferation of wing imaginal discs in Drosophila (Kawamura et al., 1999). Among those peripheral glands, feedback regulation is known only in the axis of the brain–Inka cell at ecdysis (see Chapter 3.1) and the brain–prothoracic gland axis (see Chapter 3.2). No information is available on the feedback regulation of the production and/or secretion of metabolic hormones such as hypertrehalosemic hormone and adipokinetic hormone from the corpora cardiaca (see Chapter 3.10), although they are crucially involved in the homeostasis of blood sugars and lipids, and even of allatotropin and allatostatin that control JH secretion from the corpora allata (see Chapter 3.7). In addition to feedback in the vertical axis, horizontal interactions are known between the prothoracic glands and corpora allata. Ecdysteroids and JHs interact, and we cannot study one without considering the other (Gilbert et al., 2000). Thus, the feedback regulation of the brain–prothoracic gland axis and the horizontal controls involved in ecdysteroidogenesis will be the main focus of this chapter (Figure 1).

3.8.2. Feedback Regulation of Brain–Prothoracic Gland Axis 3.8.2.1. Feedback Regulation of Brain Neurosecretory Cells by 20-Hydroxyecdysone

20-Hydroxyecdysone (20E) has been considered to be involved in the regulation of production and/or secretion of neurosecretory materials from brain NSCs based on the changes in NSC shape and stain intensity (e.g., Steel and Harmsen, 1971; Steel, 1975; Agui and Hiruma, 1977a, 1977b; Khalil et al., 1988) (see Chapter 3.11), but the evidence is only circumstantial. The weakest point of those earlier studies is the lack of knowledge of the products of the NSCs in question. Most of these studies were concerned with the four pairs of large NSCs in the pars intercerebralis of the brain, which were once believed to be prothoracicotropic hormone (PTTH) cells (Steel and Harmsen, 1971; Steel, 1975; Agui and Hiruma, 1977a) but are not now believed to be so (Agui et al., 1979; Mizoguchi et al., 1990; Gilbert et al., 2000). This indicates that ecdysteroids affect other NSCs besides the PTTH cells, although it is not truly feedback regulation. It should be noted that the nomenclature of negative and positive feedback in the earlier literatures is somewhat confusing. When 20E injections increase the stain intensity of NSCs, this was described as a positive feedback effect, although it is not certain whether the increase in stain intensity is a result of feedback inhibition of neurosecretion or activation of the gene encoding the neuropeptide or both. Both in vivo and in vitro experiments show that 20E elicits an increase in the stain intensity of most brain NSCs in Rhodnius prolixus (Steel, 1975) and Mamestra brassicae (Agui and Hiruma, 1977a). Removal of the prothoracic glands from M. brassicae larvae decreases the stain intensity and 20E injection recovers it (Agui and Hiruma, 1977b). Since the increased stain intensity may be caused by an increased production of neurosecretory materials or suppression of their release, or both, the results indicate that 20E stimulates the production of neurosecretory material (positive feedback) but suppresses its release (negative feedback) (Steel, 1975). The observations in those reports have been focused on the median NSCs, of which four pairs of the large cells are now believed to be bombyxin cells (Mizoguchi et al., 1987; Iwami et al., 1990; Moto et al., 1999) (see Chapter 3.2), suggesting that 20E affects bombyxin secretion and/ or its production. In Inka cells, that produce a peptide hormone, ecdysis triggering hormone (ETH), rising ecdysteroid levels upregulate ETH gene

Feedback Regulation of Prothoracic Gland Activity

411

Figure 1 Vertical sequence and horizontal network of regulation of prothoracic gland, based on the information obtained mainly from studies on Manduca sexta and Bombyx mori except for the growth-blocking peptide (GBP) network. Red lines indicate the vertical sequence that provides the feedback inhibition of the brain–prothoracic gland axis. Green lines indicate partly the feedback stimulation loop. Blue lines show the horizontal regulation of the prothoracic glands through the stress–GBP network (partly hypothetical). Arrowheads and ? symbols indicate stimulatory and inhibitory regulation, respectively. Shadowed area indicates that the feedback effects change along with postembryonic development: 20E inhibits the larval glands with moderate to high secretory activity, while stimulating the glands with low secretory activity as seen in early fifth instar larvae, the first day of nondiapausing pupae, and diapausing pupae of M. sexta. JH suppresses prothoracic glands before the onset of wandering, but stimulates the glands in the early pupal period. In the stress–GBP pathway, dopamine suppresses the corpus allatum until the middle of the fifth instar but not thereafter, probably due to the change in the type of dopamine receptor (Granger et al., 1996) (see Chapter 3.7). AS, allatostatin; AT, allatotropin; CA, corpus allatum; CNS, central nervous system; DDC, DOPA decarboxylase; GBP, growth-blocking peptide; NSC, neurosecretory cell; PG, prothoracic gland cell; PTSP, prothoracicostatic peptide; PTTH, prothoracicotropic hormone. See text for details.

expression, while the subsequent ecdysteroid decline triggers ETH release (Zitnan et al., 1999) (see Chapter 3.1). Although Inka cells are not NSCs, neuropeptide gene expression could be similarly upregulated in the brain NSCs, which is in accordance with the earlier observations described above. s0020

3.8.2.2. Feedback Regulation of PTTH Cells by 20-Hydroxyecdysone

There is no firm evidence to show the feedback regulation of PTTH production and its release, but 20E appears to elicit feedback control on PTTH cells, as seen in the diapausing pupal brain of M. brassicae (Agui and Hiruma, 1977a) (see Chapter 3.12). Diapausing pupal brains were incubated with 20E for 2 days followed by incubation in fresh

medium for another 2 days, and then inactive diapausing pupal prothoracic glands were placed in the latter conditioned medium. When ecdysteroid activity in the final medium was assessed biologically by adding a piece of integument, apolysis (secretion of new cuticles from the underlying epidermal cells) was induced, indicating that the conditioned medium in which prothoracic glands from diapausing pupae had been added did contain ecdysteroids. This suggested that the 20E in the first medium might have caused the brain to release PTTH. They also showed increases in the stain intensity of all median and posterior brain NSCs, but not in the lateral NSCs. The intensity of the latter actually decreased after incubation of the brain with 20E for 2 days, but it is not obvious whether these lateral NSCs are the PTTH cells that have been described as

412 Feedback Regulation of Prothoracic Gland Activity

being in the lateral region of other Lepidoptera (Agui et al., 1979; Mizoguchi et al., 1990; Gilbert et al., 2000) (see Chapter 3.2) since there are at least three types of NSCs of similar size in the lateral region. Nevertheless, the data suggest that 20E stimulates PTTH release at a moderate concentration while suppressing its release at a high concentration (Agui and Hiruma, 1977a). Circumstantial evidence for the above hypothesis also comes from comparing the changes in hemolymph ecdysteroid and PTTH concentrations in the fourth and fifth larval instars and the pupal period of Bombyx mori (Mizoguchi et al., 2001, 2002). In the fourth instar, the PTTH titer begins to decline as the ecdysteroid titer peaks. Brain extirpation or neck ligation has been utilized to estimate the time after which the brain is not required for inducing the following larval ecdysis, referred to as the critical period, and thereby the time when sufficient PTTH is secreted (Truman, 1972). This time is approximately 14 h prior to the peak of the ecdysteroid titer in Manduca (Bollenbacher et al., 1987) and 12 h in Bombyx fourth instar larvae (Sakurai, 1983). This time is well correlated with the change in PTTH titer. Nevertheless, the question remains concerning the extremely low PTTH titer as low as 18 pg ml1 hemolymph in Bombyx (Mizoguchi et al., 2001), which is an order of magnitude lower than the critical concentration (0.2–0.3 ng ml1) to stimulate the prothoracic glands in vitro (Kataoka et al., 1987; Kiriishi et al., 1992; Hua et al., 1999). Accordingly, it is not conclusive at present if the measured PTTH titer in the fourth instar larvae reflects the actual PTTH concentration or if there are other factors involved in the upregulation of prothoracic gland activity, although brain extracts effectively induce a larval–larval molt in neck ligated larvae (Suzuki and Ishizaki, 1986). In Manduca, PTTH titers in the fourth instar larval hemolymph were measured by an indirect bioassay method using paired prothoracic glands (Bollenbacher et al., 1987). The peak titer of PTTH was found to be 0.2 units of PTTH, just enough to activate the glands, and occurred 9 h prior to the ecdysteroid peak (Bollenbacher et al., 1987). When the ecdysteroid titer peaks, the PTTH titer has decreased, indicating the feedback inhibition of PTTH synthesis and/or release by 20E. At that stage any ecdysone formed would be promptly converted to 20E (Kamimura and Kiuchi, 2002). In the fifth instar, the PTTH titer declines sharply in concurrence with the beginning of the increase in ecdysteroid titer approximately 12 h later (Mizoguchi et al., 2002). This supports the idea of feedback inhibition of PTTH release by high ecdysteroid titers.

A similar observation was made for the pupal period (Mizoguchi et al., 2002). In Bombyx pupae, the PTTH titer peaks on day 1.25 and declines after considerable fluctuations. The peak ecdysteroid titer occurs 1 day later, although both peaks are broad and overlap. The ecdysteroid titer was measured by radioimmunoassay (RIA) of crude extracts of hemolymph and the changes in individual ecdysteroids were determined by a combination of high-performance liquid chromatography (HPLC) separation and RIA measurement, a paradigm that showed that the 20E peak follows the ecdysone peak by 3 days in Manduca (Warren and Gilbert, 1986). Taking into consideration that pupal–adult development takes 9 days in Bombyx, which was the object of the studies by Mizoguchi et al. (2001), while the comparable period in Manduca is about 20 days, the 20E peak in Bombyx pupae should appear at least 1 day after the measured ecdysteroid peak. In that case, the PTTH titer would decline when the 20E titer has peaked. Accordingly, 20E may downregulate PTTH secretion and/or synthesis throughout the postembryonic development, except in the case of diapausing animals. It has not been shown unequivocally that 20E exerts feedback regulation on PTTH production. At each of the developmental stages of Bombyx, a high ecdysteroid titer does not elicit an accumulation of PTTH in the brain (Mizoguchi et al., 2001) nor in the PTTH cell axon ending of the corpus allatum of Manduca (Dai et al., 1995), a neurohemal organ for PTTH, in contrast to brain NSCs indicated by the above-mentioned earlier observations (Steel, 1975; Agui and Hiruma, 1977a). It is conceivable that once 20E elicits its inhibitory effects on the PTTH cells at its high concentration at the end of the fourth and fifth instars, 20E may not only suppress PTTH secretion but also downregulate the PTTH gene. In the hypothalamic– pituitary–adrenal or –thyroid axes in vertebrates, corticosteroid or thyroid hormone downregulates the genes of tropic components in the hypothalamic NSCs and pituitary cells. The hypothalamic TRH and pituitary TSH subunit genes are downregulated by the active form of thyroid hormone, triiodothyronine (T3) (Shibusawa et al., 2003). There is no indication if such is also the case in PTTH gene regulation, since we know nothing about this gene, but 20E could exert its effect by downregulating the PTTH gene. Another interpretation is suggested by the study of deiodinase activity in the thyrotrophs of the anterior pituitary gland (Huang et al., 2001; Schneider et al., 2001). When TSH genes are suppressed strongly by thyroid hormone, type II iodothyronine deiodinase (D2) activity

Feedback Regulation of Prothoracic Gland Activity

increases in the thyrotrophs to promote the conversion of T4 to T3, the latter being stronger in its ability to bind to the thyroid hormone receptor and thereby promote binding to trans-regulatory elements of the TRH and TSH genes (Shibusawa et al., 2003). Such locally synthesized T3 acts as a negative feedback factor in suppressing the TSH gene, and therefore the localized upregulation of D2 activity may be the first step in the negative feedback loop (Huang et al., 2001). Although it is not known if circulating 20E in insect hemolymph is positively released from the midgut and fat body, major tissues expressing 20-monooxygenase activity (Smith et al., 1983), i.e., the enzyme that mediates the conversion of ecdysone to 20E, or only leaks from peripheral tissues. It might be of interest to measure this enzyme activity in PTTH cells when they are presumably negatively controlled by 20E. If the 20-monooxygenase is upregulated locally in the PTTH cells, feedback inhibition of the PTTH gene may occur earlier than the time when a 20E increase occurs in the hemolymph. Since the gene encoding the 20-monooxygenase is now known (see Chapter 3.3), one could use in situ hybridization studies to quantify that gene’s transcripts in the PTTH cells. In addition to PTTH, prothoracicostatic peptide (PTSP) appears to be present in Bombyx fifth instar larvae, although it is identical to the Manduca myoinhibitory peptide (Mas-MIP I) (Hua et al., 1999). PTSP does not alter the basal secretory activity of prothoracic glands in vitro but suppresses the PTTH-stimulated glands, presumably by inhibiting PTTH action (Hua et al., 1999) by blocking the calcium channel that is tightly associated with the earliest step of the PTTH signaling pathway (Dedos and Birkenbeil, 2003). PTSP has been isolated from the Bombyx brain, and therefore the site of synthesis is probably in the brain, but nothing is known about the location of the PTSP cells or the effects of 20E on its production and/or secretion.

3.8.3. Feedback Regulation of the Prothoracic Gland by Ecdysteroid A decrease in the hemolymph ecdysteroid titer may be as important as its increase in various aspects of insect development. At the larval–larval molt, the DOPA decarboxylase (DDC) gene in the epidermal cells is activated in response to a decreasing ecdysteroid titer (Hiruma et al., 1995). DDC mediates the conversion of DOPA to dopamine, a precursor of DOPA quinone that mediates screlotization by chemically bridging cuticular proteins (see Chapters 4.2 and 4.4) and which also serves as an

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intermediate in melanin production for melanization. Therefore, an artificial increase in ecdysteroid concentration delays the melanization process. Similarly, adult eclosion is delayed by 20E injection (see Chapter 3.1). In spite of the recognition of the importance of feedback regulation of prothoracic gland secretory activity, only six papers on this subject have been published in the last 50 years (Williams, 1952; Siew and Gilbert, 1971; Beydon and Lafont, 1983; Bodnaryk, 1986; Sakurai and Williams, 1989; Song and Gilbert, 1998; review: Gilbert et al., 1997). The next section is based on those publications. 3.8.3.1. Feedback Activation of Ecdysteroid Production

The first paper that indicated the feedback activation of prothoracic glands appeared in 1952, although the term ‘‘feedback’’ was not used. Rather, it was stated that ‘‘prothoracic glands can be triggered also by the prothoracic gland hormone itself’’ in the famous experiments involving the parabiosis of eight brainless diapausing pupae and chained pupal abdomens of Hyalophora cecropia (Williams, 1952). This study doubtlessly describes the positive feedback regulation of prothoracic gland activity. There are three reports to support feedback activation in Manduca (Sakurai and Williams, 1989), the bertha armyworm Mamestra configurata (Bodnaryk, 1986), and Bombyx (Sakurai and Imokawa, 1988) although Bodnaryk does not mention positive feedback. The prothoracic glands of day 2 fifth instar Manduca larvae were inserted into diapausing pupae that had had their brains removed, and gland activity was then measured in vitro every day after implantation. In such pupal preparations, the implants exhibited periodic changes in secretory activity, as the peaks of 50–60 ng per gland appeared 4, 7, and 12 days after the implantation. Each peak was separated by a nadir of activity of approximately 20 ng per gland. The same was also the case in pupal preparations lacking both brain and prothoracic glands. The implanted day 2 glands may be stimulated by ecdysteroid they synthesize. When the activity attains a considerably higher level than a threshold in the host pupae, the glands are inhibited. Once the gland activity decreases to a low level, they are again capable of being activated by their ecdysteroids or those of the host prothoracic glands that may have been activated by the ecdysteroid produced by the implanted glands. These in vivo observations indicate the presence of feedback activation. However, the glands of feeding larvae are barely activated by either ecdysone or 20E in vitro. By contrast, in vitro administration of 20E stimulates diapausing pupal

414 Feedback Regulation of Prothoracic Gland Activity

glands as well as day 1 nondiapausing pupal glands (Sakurai and Williams, 1989). A similar autonomous change in the gland activity is observed in Bombyx fifth instar larvae (Sakurai and Imokawa, 1988). Larvae lacking the retrocerebral complex (brain–corpora cardiaca–corpora allata) are capable of gut purge followed by pupation if the surgical operation was done on day 4 of the instar, 2 to 3 days before gut purge. After the operation, the gland activity gradually increases and gut purge is induced. Subsequently, the activity sharply decreases to its lowest level before pupation. Although in vitro challenge by ecdysteroids failed to activate glands of feeding Manduca larvae, the composite data indicate that 20E causes stimulation of ecdysteroid production of glands when their secretory activity is as low as those of diapausing pupae, but the feedback activation of young last instar larval glands of relatively low secretory activity is conjectural. In M. configurata pupae, injection of 20E (500 ng g1 fresh weight) into postdiapause, pupae 1 day after transfer to 20  C from 6 to 8 months of storage at 0  C accelerates the increase in ecdysone concentration in the hemolymph in about 1 day and in the 20E concentration in about 2 days. In contrast, a higher 20E dose, as much as 2500 ng g1, suppressed the occurrence of both ecdysteroid peaks (Bodnaryk, 1986), as will be discussed below (see Section 3.8.3.2). In postdiapause pupae, ecdysone and 20E titers begin to increase 4 and 7 days after being transferred to 20  C, showing that the glands 1 day after the transfer are apparently not yet activated, and therefore, indicates that the exogenous 20E stimulated the glands having very low or no secretory activity, as in the H. cecropia and Manduca diapausing pupal glands. In diapausing pupae, either photoperiod induced or artificially induced by brain removal from nondiapausing pupae at pupation, adult development initiates spontaneously if it is not a deep diapause, as for the univoltine H. cecropia. In brainless pupae, there is no PTTH source, but in such Manduca pupae, ecdysone is the major ecdysteroid and [3H]-cholesterol is incorporated into ecdysone at the beginning of spontaneous adult development, indicating that the prothoracic glands were activated in those pseudodiapausing pupae (Sakurai et al., 1991). Since they lack a brain, the prothoracic glands may be activated by signals other than PTTH, possibly by the positive feedback of ecdysteroid that the prothoracic glands secreted. Although feedback activation has been recorded for pupal prothoracic glands, there is no evidence that such activation is responsible for the increase in ecdysteroid titer that triggers pupal–adult development.

3.8.3.2. Feedback Inhibition

Siew and Gilbert (1971) first showed the negative feedback regulation of prothoracic gland activity. This is more easily explainable than that discussed above, i.e., to downregulate the glands once they have secreted enough ecdysteroid to initiate the molting process. Later, feedback inhibition of the prothoracic glands was described in the large cabbage butterfly Pieris brassicae (Beydon and Lafont, 1983), M. configurata (Bodnaryk, 1986), and M. sexta (Sakurai and Williams, 1989; Song and Gilbert, 1998). In Pieris pupae, the inhibition of ecdysteroid biosynthesis by exogenous 20E is dose dependent (Beydon and Lafont, 1983). In Manduca, when prothoracic glands at peak activity 2 days after the onset of wandering were implanted into pupae from which the brain–corpora cardiaca– corpora allata complex and prothoracic glands had been removed, the 3-dehydroecdysone (see Chapter 3.3) secretory activity of the implanted glands decreased at the same rate as those in intact larvae. Similar inhibition is elicited by 20E in vitro, indicating that feedback inhibition solely accounts for the decrease of secretory activity after the peak of activity that occurs prior to pupation (Sakurai and Williams, 1989). Feedback inhibition by 20E was demonstrated clearly using the prothoracic glands of day 5 Manduca larvae (Song and Gilbert, 1998). The glands were incubated in the presence or absence of 20E for various time periods and then incubated in medium alone for a further 1 h to assess the basal secretory activity (Figure 2). The basal level of ecdysteroid production was markedly inhibited by the 1 h incubation with 20E to approximately 25% of the control. When these glands were challenged with PTTH after exposure to 20E for 24 h, the enhanced level of ecdysteroidogenesis was far less than that of the control glands. Comparison of basal ecdysteroidogenesis with PTTH-stimulated activity after gland incubation for 6 h with 20E, showed that the glands were still capable of responding to PTTH, although the basal level was very low due to the inhibition by 20E. By contrast, after 24 h incubation in medium alone, the activation ratio (amount of ecdysteroids produced by one of a paired set of glands incubated with PTTH/amount of ecdysteroids produced by the contralateral gland incubated in medium alone) (Agui et al., 1979) was 11 while it was 2.5 after the incubation with 20E. In addition, the basal secretory activity after the incubation with 20E for 6 h was similar to that after incubation in medium alone for 24 h, but the activation ratio for the former glands was 4.5 whereas that for the latter glands

Feedback Regulation of Prothoracic Gland Activity

Figure 2 Inhibition by 20E of basal secretory activity and the responsiveness to PTTH of the prothoracic glands. One of a paired prothoracic glands of day 5 fifth instar Manduca larvae were incubated with 10 mM 20E (red and pink bars) or in medium alone (green and light green bars) for an indicated period and then incubated for another 1 h in a medium containing PTTH (red and green) or in medium alone (light green and pink) for assessing the basal secretory activity and the gland responsiveness to PTTH. Inset table: activation ratio (amount of ecdysteroids produced in the presence of PTTH/amount of ecdysteroids produced in medium alone) calculated from the paired values in the figure. Note that 20E suppresses the basal secretory activity at 1 h of incubation with 20E (light green vs. pink), while hardly affecting the PTTH-stimulated secretory activity (green vs. red). The latter is suppressed by a 6 h incubation with 20E. (Modified from Song, Q., Gilbert, L.I., 1998. Alterations in ultraspiracle (USP) content and phosphorylation state accompany feedback regulation of ecdysone synthesis in the insect prothoracic gland. Insect Biochem. Mol. Biol. 28, 849–860.)

was 11, indicating that the 6 h incubation with 20E partially suppressed the gland responsiveness to PTTH. Thus, the prothoracic glands retained their competence to PTTH after a short incubation with 20E, suggesting that 20E simply suppressed the ecdysteroidogenic machinery, and that an exposure to 20E for a considerably longer period inhibited PTTH-stimulated ecdysteroidogenesis as well. Feedback inhibition is known in crustaceans (see Chapter 3.16). The secretory activity of the Y-organs

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of the crayfish Orconectes limosus was greatly inhibited within 1 h after a single injection of 20E and slightly, but significantly, by a 1 h incubation with an ecdysteroid antagonist, RH-5849 (Dell et al., 1999). This result is very similar to the in vitro feedback inhibition of Manduca prothoracic glands. They were markedly inhibited within 1 h by incubation with 20E (Song and Gilbert, 1998) or RH-5849 (Sakurai and Gilbert, unpublished data). Therefore, the feedback inhibition of the ecdysteroidogenic organs could be common to arthropods. Ecdysone exerts feedback inhibition of Manduca day 3 fifth instar prothoracic glands in vitro, although the effect is faint (Sakurai and Williams, 1989). When the Manduca glands were inhibited by ecdysteroids (see Section 3.8.5.1), USP-1 and USP-2 proteins appeared in complementary patterns with USP-2 being predominant (see Chapter 3.5). Both suppression of USP-1 protein synthesis and simultaneous up-regulation of USP-2 synthesis are brought about by ecdysone and 20E. The effective concentration of ecdysone is, however, 100-fold as much as 20E (Song and Gilbert, 1998). This weak effect of ecdysone is consistent with the Kd of ecdysone for the nuclear receptor complex, which is lower than that of 20E by a factor of more than 100 (Cherbas et al., 1980) (see Chapters 3.5 and 3.6). Since the prothoracic glands express no 20-monooxygenase activity mediating the conversion of ecdysone to 20E (Warren et al., 1988), the result still indicates a weak action of ecdysone. Although ecdysone metabolites have been proposed to have morphogenetic roles of their own (Gilbert et al., 2000; Gilbert, personal communication), no information is available on the possibility that 20,26-dihydroxyecdysone, the immediate metabolite of 20E, exerts feedback inhibition on prothoracic glands. 3.8.3.3. Complete and Incomplete Inactivation

There are two types of suppression of prothoracic gland secretory activity. One is incomplete shutdown, as seen at the ecdysis from the third (second penultimate) to the fourth (penultimate) instar and also at the end of the fifth (last) instar associated with the climax of pupal metamorphosis. In those periods, the gland activity declines greatly but still remains at a low level as the glands are still capable of secreting considerable amounts of ecdysone when examined in vitro (Okuda et al., 1985; Bollenbacher et al., 1987). In addition, they retain competence to respond to PTTH (Okuda et al., 1985; Gu et al., 2000; Takaki and Sakurai, 2003). By contrast, at the time of ecdysis from the penultimate to the last larval instar in Bombyx, gland activity is completely shut down for a period of 2–3 days (Okuda et al.,

416 Feedback Regulation of Prothoracic Gland Activity

1985; Gu et al., 1997). In Manduca, the in vitro secretory activity of the glands on the second day of the fifth instar is 0.01–0.03 ng h1 per gland (Smith and Pasquarello, 1989; Rybczynski and Gilbert, 1994) (see Chapter 3.2), indicating that activity is at an almost negligible level. In this stage in Bombyx, the glands are insensitive to PTTH. It is not clear if this insensitivity is due to the disappearance of the PTTH receptor from the gland cell membranes or a deficiency of one or more factors involved in the PTTH signaling. Whatever the cause, JH is involved in maintaining this inactivity. When the corpora allata are removed from newly molted Bombyx fifth instar larvae, the basal secretory activity of their glands reappears 9 h after the allatectomy. Three hours before this time, the glands exhibit a normal response to PTTH (Sakurai et al., 1989a). This indicates that all the apparatus for ecdysteroidogenesis has been recovered before initiating spontaneous ecdysone secretion, and JH acts to maintain the suppressed condition (Takaki and Sakurai, 2003). Thus, feedback inhibition late in the penultimate instar may affect some steps of ecdysteroidogenesis and such a deficiency is maintained by JH. However, there is no information about whether the PTTH receptor itself has disappeared, either due to repression of its gene expression or it is simply sequestered away by the involution of the plasma membrane into the inside of the cells. Because dibutyryl cAMP and IBMX, a phosphodiesterase inhibitor, have lost their ability to enhance ecdysteroidogenesis at this stage (Gu et al., 2000; Takaki and Sakurai, 2003), a simple deficiency of PTTH receptors can hardly account for the gland’s inactivity.

3.8.4. Mechanisms of Decrease in Hemolymph Ecdysteroid Titer: Biochemical Aspects s0050

3.8.4.1. Inactivation of 20-Hydroxyecdysone

There are at least two possible reasons for the decrease in the hormone titer, i.e., an increase in inactivation and excretion, and/or a decrease in hormone production. When ecdysone is injected into the body cavity of either a fifth (last) instar larvae or adult of Locusta migratoria, the ecdysteroid concentration at first increases, but promptly declines (Hoffmann et al., 1974). The fate of the injected ecdysone is stage specific. On day 7 of the 11 day long instar when the hemolymph ecdysteroid titer is at its highest, ecdysone is promptly converted to 20E within 2 h and then is slowly excreted with the feces. In contrast, in either early or late fifth instar larvae or adults, the injected ecdysone is excreted promptly

into the feces as is, or after conversion to inactivation products. This indicates that at the time of peak ecdysteroid titer, feedback inhibition by 20E may act to increase the excretion process, since at this time, 20-monooxygenase activity is maximal while the excretion rate is at its lowest. This results in the accumulation of 20E in the hemolymph. In the metabolism of ecdysteroids, there are two immediate inactivation steps, epimerization and hydroxylation (see Chapter 3.3). Epimerization occurs at C3 to form 3-epiecdysteroid via an intermediate, 3-dehydroecdysteroid (Webb et al., 1995). Since the hemolymph contains a high level 3-dehydroecdysone 3b-reductase activity (Sakurai et al., 1989b; Kiriishi et al., 1990), it is not possible to reduce 3-dehydroecdysteroid to 3-epiecdysteroid in hemolymph by the 3a-ketoreductase. Accordingly, inactivation of ecdysteroid through 3-epimerization is restricted only to tissues. Another immediate inactivation reaction of 20E, and probably the most important for 20E metabolism, is brought about by hydroxylation at the terminal carbon of the side chain to form 20,26-dihydroxyecdysone. This is further oxidized to the 26-oic acid and exhibits no biological activity. In the midgut cytosolic fraction prepared from feeding larvae of Manduca, the 26-hydroxylase is induced by 20E through the upregulation of its gene (Williams et al., 1997). Although we have no information on 20,26-dihydroxyecdysone concentrations in larval hemolymph, it is highly possible that it achieves a maximum following that of 20E, similar to what occurs in Manduca pharate adults. Since the 26-hydroxylase is induced by 20E during the larval period, it would be of interest to examine the feedback effects of 20,26-dihydroxyecdysone on ecdysteroidogenesis. In Manduca pharate adults, the 20,26-dihydroxyecdysone concentration increases after the peak in the 20E titer, and becomes the sole free ecdysteroid metabolite among the various radioimmunoreactive steroids in the hemolymph at this stage (Warren and Gilbert, 1986). Accordingly, the upregulation of the 26-hydroxylase gene by 20E may be the ultimate cause of the decrease in the hemolymph 20E titer during the pupal– adult development. However, this may not be the sole reason for the decrease in 20E titer, since at this period, ecdysone secretion has almost ceased due to the degeneration of prothoracic glands, the result of programmed cell death induced by 20E (Dai and Gilbert, 1997, 1998). 3.8.4.2. Suppression of Ecdysteroidogenesis

There may be at least three different targets for the negative feedback induced by 20E in prothoracic gland cells: the enzymes of the ecdysone biosynthetic

Feedback Regulation of Prothoracic Gland Activity

pathway, the hypothetical system for the subcellular movement of intermediary ecdysteroid metabolites, and the signaling pathway for PTTH (Figure 3). Ecdysone synthesis is a complicated pathway and our knowledge is still rudimentary. We have extensive knowledge of only some of the steps occurring in the prothoracic glands (Gilbert et al., 2002) (see Chapter 3.3). The first reaction is the dehydrogenation of cholesterol to 7-dehydrocholesterol by a P450 (CYP) enzyme in the endoplasmic reticulum (ER) (Grieneisen et al., 1993; Warren and Gilbert, 1996). It is followed by a series of complex and uncharacterized oxidation reactions occurring in a location that is presently unknown, although it has long been thought to be the mitochondria in analogy with mammalian steroidogenesis (see below). The product of these reactions, an ecdysteroid precursor, is then hydroxylated at C25 by another CYP enzyme present in the ER (Kappler et al., 1988), followed by

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additional hydroxylations at C22 and finally C2, both catalyzed by different CYP enzymes present in the mitochondria (Warren et al., 2002) (see Chapter 3.3). The product, either 3DE and/or E (Warren et al., 1988), is then secreted from the prothoracic gland cell by another unknown process and subsequently is taken up into peripheral tissues and converted to 20E by ecdysone-20-monooxygenase, another CYP enzyme variously localized in both the ER and mitochondria (Petryk et al., 2003). 20E possibly affects one or more of these enzymes involved in the biosynthesis of ecdysone, but the effects of 20E on P450 activities have not yet been investigated. Involvement of both the ER and mitochondria in ecdysone biosynthesis indicates that both cholesterol and ecdysone precursors must necessarily be conveyed between critical subcellular domains, including but not limited to the ER and the mitochondria, during ecdysone biosynthesis. Some of

Figure 3 Feedback regulation of ecdysteroidogenesis in the prothoracic gland cell. Red lines indicate the ultra-short-loop feedback inhibition, which has been so far revealed. Arrowheads and ? symbols indicate activation and inhibition, respectively. Note that 20E binds to USP-2 and activates EcR/USP-2 though the responsible isoform of EcR has not yet been identified. Then the nuclear receptor heterodimer acts as a transcription activator and the resulting proteins may suppress the genes whose products are involved in ecdysteroidogenesis (Song and Gilbert, 1998). See Chapter 3.2; for details of the PTTH signaling pathway and see Chapter 3.3 for the ecdysone synthesis pathway. The membrane receptor for 20E is hypothetical. AC, adenylyl cyclase; cAMP, cyclic AMP; CaMK, Ca2þ/calmodulin-dependent protein kinase; 3DE, 3-dehydroecdysone; E, ecdysone; ER, endoplasmic reticulum; MAPK, mitogen activated protein kinase; MT, mitochondrion; PKA, cAMP-dependent protein kinase; PTSP, prothoracicostatic peptide; PTTH, prothoracicotropic hormone; S6, ribosomal protein S6; S6K, S6 kinase.

418 Feedback Regulation of Prothoracic Gland Activity

these sterol translocations may be rate limiting in Manduca and Drosophila ecdysone synthesis (Warren and Gilbert, 1996; Watson et al., 1996), and therefore, a possible target of negative feedback regulation. In mammalian steroidogenic cells, the primary substrate, cholesterol, is transported from its intracellular reservoirs, the cell membrane and/or the ER, first to the outer mitochondrial membrane and then to the CYP enzyme present in the inner mitochondrial membrane (P450ssc), perhaps with the aid of the cytoskeleton and/or various ‘‘translocation’’ factors, and these processes are accelerated upon stimulation by a tropic hormone (Hall, 1995, 1997). In Manduca prothoracic glands, PTTH stimulates the synthesis of b-tubulin, which is rapid and transitory, suggesting a direct role of microtubules in the upregulation of ecdysteroidogenesis by PTTH (Rybczynski and Gilbert, 1995a) (see Chapter 3.2). Indeed, microtubule-disrupting drugs inhibit PTTH-stimulated ecdysteroidogenesis in the pupal prothoracic glands (Watson et al., 1996). Although the drugs do not inhibit the basal secretory activity of pupal glands, nor PTTH-stimulated larval glands (Gilbert et al., 2002), the turnover of cytoskeletal proteins is a possible target of acute feedback inhibition, since vertebrate and insect steroidogenic cells are surprisingly homologous in the mechanisms that regulate the changes in steroidogenesis (Gilbert et al., 2002). A third possibility includes factors that modulate or control, directly or indirectly, the PTTH signaling pathway (see Chapter 3.2). PTTH stimulation is known to require de novo gene expression and protein synthesis (Keightley et al., 1990; Rybczynski and Gilbert, 1994), and the latter may be mediated by S6 kinase (S6K) which mediates ribosomal S6 phosphorylation (Song and Gilbert, 1994, 1995, 1997). Circumstantial evidence deduced from time-course studies indeed shows that PTTH-stimulated S6 phosphorylation is involved in the upregulation of ecdysteroidogenesis. Since S6K activates several transcription factors, including c-Jun and Atf-2, PTTH signaling could involve activation of the transcription factors. Interestingly, the dephosphorylation of S6 protein occurs in temporal synchrony with the decline in ecdysone biosynthesis (Song and Gilbert, 1995), indicating that the feedback inhibition of ecdysteroidogenesis does not necessarily involve the direct inhibition of the enzymes in the ecdysone biosynthetic pathway. Rather, the regulation of the synthesis of proteins that control such enzyme activities could be the primary target of negative feedback regulation by 20E. In PTTH-stimulated prothoracic gland cells, hsp70 synthesis is also upregulated (Rybczynski and Gilbert,

1995b, 2000), implying its participation in feedback control of ecdysteroidogenesis by the modulation of the assembly of the ecdysteroid receptor complex. In this regard, USP proteins are suggested to exert important roles in the negative feedback (Gilbert et al., 1997) (see Section 3.8.5). Although not yet identified, molecular chaperones besides hsp70 could also be involved in modulating the rate of ecdysteroidogenesis by assisting the movement into the mitochondria of catalytic enzymes synthesized in the cytosol.

3.8.5. Molecular Mechanism of Feedback Inhibition 3.8.5.1. Feedback Mediated by the EcR/USP Heterodimer

In the feedback inhibition of prothoracic glands, the primary action of 20E is most likely mediated by a heterodimeric nuclear receptor composed of the ecdysone receptor (EcR) and Ultraspiracle (USP) (see Chapters 3.5 and 3.6). In this interaction, the USP isoform is of primary importance (Gilbert et al., 1997; Song and Gilbert, 1998) (Figure 3). Developmental profiles of two USP isoforms in the prothoracic glands of Manduca last instar larvae revealed that USP-1 and USP-2 are expressed in complementary patterns. At first, USP-1 is predominant while USP-2 is barely detectable until day 6 at which time the hemolymph ecdysteroid level only begins to rise. After day 6, USP-2 is upregulated and expressed maximally on days 7–9, i.e., during the peak in the ecdysteroid titer, while USP-1 is simultaneously downregulated to almost undetectable levels. The net result is that USP-2 comprises about 85% of the total USP proteins (Gilbert et al., 1997). The downregulation of USP-1 and simultaneous upregulation of USP-2, both being elicited by 20E, may alter a set of genes that ultimately leads to downregulation of ecdysteroidogenesis. In vitro incubation with 20E inhibits gland activity (Figure 2). Incubation of day 5 prothoracic glands with 20E reduced the basal secretory activity in 6 h to one-sixth that of the control glands incubated in medium alone. During the 6 h incubation with 20E, USP-1 is downregulated while USP-2 is upregulated. In day 7 glands, only the EcR/ USP-2 heterodimer is present, but not EcR/USP-1, indicating that USP-2 becomes the predominant component of the receptor complex in response to a high ecdysteroid titer, forming EcR/USP-2, presumed to be functional in feedback inhibition (Song and Gilbert, 1998). After the binding of 20E to EcR, the functional heterodimer probably leads to the downregulation of the expression of one or more critical

Feedback Regulation of Prothoracic Gland Activity

components of the ecdysteroidogenic pathway (Gilbert et al., 1997). 20E is more effective than ecdysone by a factor of 100 in eliciting the expression of USP-2 in day 5 prothoracic glands. This difference in activity coincides well with the difference in binding affinities of these two ecdysteroids for the EcR/USP complex (Cherbas et al., 1980; Minakuchi et al., 2003) (see Chapter 3.5). Such a difference is reflected in the activities of ecdysone and 20E in the in vitro feedback inhibition of the prothoracic glands (Sakurai and Williams, 1989). Two EcR isoforms, EcR-A and EcR-B1, have been identified in Lepidoptera. However, it remains to be seen which isoform participates in forming the functional EcR/USP-2 heterodimer and whether the EcR/ USP-2 heterodimer is the primary transcriptional factor providing for the downregulation of USP-1. Although incubation of day 5 Manduca prothoracic glands with 20E for 6 h markedly reduces the basal secretory activity, they still retain their responsiveness to PTTH, e.g., as PTTH elicits an activation ratio of 5 (Figure 2). This activation ratio, however, decreases from approximately 15 after 1 h incubation with 20E, to 5 after 6 h, indicating that 20E does affect the PTTH signaling pathway. The complementary patterns with USP-2 being predominant during the incubation with 20E could cause the loss of sensitivity to PTTH (Song and Gilbert, 1998), and thereby act to sustain the inhibited state of ecdysteroidogenesis. It is of interest that the developmental profiles of individual USP isoform expression in epidermis are quite similar to those in the prothoracic glands (Hiruma et al., 1999). USP-1 mRNA levels are high during the feeding period, but decline to low levels shortly after the onset of the wandering stage when the hemolymph ecdysteroid titer begins to increase. In contrast, the USP-2 mRNA level remains low until the increase in ecdysteroid titer, after which its expression is enhanced. A similar control over the expression of USP isoforms by 20E is reported in the adult mosquito Aedes aegypti (Wang et al., 2000). After a blood meal, the fat body initiates the production of yolk protein (vitellogenin), needed for egg maturation, in response to 20E originating from the ecdysone secreted by the ovaries that are stimulated by the blood meal (see Chapter 3.9). USP-A (A. aegypti homolog of USP-1) is downregulated at the time of the increase in 20E titer, while USP-B (homolog of USP-2) is upregulated. Incubation of the fat body with 20E results in the upregulation of USP-B while USP-A transcription is downregulated. EcR/USP-B elicits a stronger transactivating activity on the vitellogenin gene than does EcR/USP-A. The function of the receptor complex, of course, varies

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among the tissues that exhibit tissue-specific responses to 20E, e.g., the feedback inhibition of ecdysteroidogenesis in the prothoracic glands, the triggering of pupal differentiation in the epidermis, and yolk protein expression in the ovaries. However, the fundamental molecular mechanisms underlying the control of gene expression by EcR/USP could be similar in individual tissues. We presently have no information to speculate about whether feedback inhibition affects one or more of the enzyme activities involved in the ecdysone biosynthetic pathway. A decrease in ecdysone production may be as important as an increase with respect to the regulation of insect growth and development, but little is known at present about feedback regulation at either the biochemical or molecular levels. 3.8.5.2. Acute and Delayed Inhibition

Incubation of day 5 fifth instar Manduca prothoracic glands with 20E for 1 h strongly suppresses the basal secretory activity to about 20% of the control, but the glands retain their competence to respond to PTTH (Song and Gilbert, 1998) (Figure 2). Similarly, RH-5849 significantly suppresses the secretory activity within 10 min while retaining sensitivity to PTTH (Sakurai and Gilbert, unpublished data). Acute inhibition is also reported in the crayfish Orconectes limosus, in which the Y-organ is the ecdysteroidogenic gland, and 20E injection to the crayfish larvae inhibits the gland activity. Ecdysteroid release from intermolt Y-organs is reduced sixfold within 1 h after 20E injection (Dell et al., 1999). In Manduca prothoracic glands, the complementary pattern of USP-1 and USP-2 expression appears only after a 6 h of incubation with 20E, but nothing is observed after only 1 h (Song and Gilbert, 1998). In an injection study in Orconectes, the degree of inhibition caused by RH-5849 at its minimum effective dose of 5 mg is less than that resulting from 0.25 mg 20E. Thus, RH-5849 is at least 20 times less effective than 20E. The binding affinity of 20E for the Chironomus tentans EcR/USP complex is in the range of 107 M (Kd ¼ 230 nM), and is similar to that of RH-5849 (Kd ¼ 278 nM) (Smagghe et al., 2002) (see Chapter 3.5). The binding affinities of 20E and RH-5849 to the in vitro translated EcR/ USP complex of Chilo suppressalis shows that 20E affinity is only threefold higher than RH-5849 (Minakuchi et al., 2003). However, it is not obvious whether the direct application of this Kd data to Orconectes is appropriate, since the binding affinity of some ecdysteroid agonists differs greatly in different orders of insects, e.g., the binding affinity of RH-2485, another potent nonsteroidal ecdysone

420 Feedback Regulation of Prothoracic Gland Activity

agonist, for crude nuclear extracts is 70-fold higher than that of 20E in the moth Plodia interpunctella, while in the beetle Anthonomus grandis, the former value is less than the latter by a factor of 50 (Dhadialla and Tzertzinis, 1997). Nevertheless, in Chironomus, Chilo, and Orconectes, in spite of the similarity in the affinities of 20E and RH-5849 to the EcR/USP complex, 20E is 20-fold more effective than RH-5849 in inhibiting ecdysone secretion, indicating that the acute inhibition may not be due to a genomic action of 20E. Hence, the acute inhibition of basal secretory activity indicates that the rapid feedback inhibition of ecdysteroid secretion might be mediated by a nongenomic pathway, rather than a conventional genomic pathway. The acute inhibition does not, however, suppress the prothoracic gland responsiveness to PTTH (Song and Gilbert, 1998). The above discussion gives rise to a more complex model for the role of feedback inhibition. There may be two different signaling pathways underlying the feedback inhibition; one is acute and the other is delayed. Acute inhibition may result in the rapid decline of ecdysone secretion after the physiological peak concentration of 20E is attained in the hemolymph and is probably mediated by a nongenomic action of 20E. After this acute inhibition, a second genomic action via the EcR/USP-2 heterodimer ensures that the glands will not produce ecdysone again once the 20E titer declines below the threshold level for the acute feedback inhibition. Thereby, the glands are maintained inactive until the next PTTH stimulation. During this period with low secretory activity, developmental events are allowed to occur in response to a declining ecdysteroid concentration.

3.8.6. Horizontal Control of the Prothoracic Gland 20E also affects CA activity while JH alters prothoracic gland ecdysteroidogenic activity (see Chapter 3.7). JH is believed to elicit the control of prothoracic gland activity in two ways: one through the control of PTTH release and the other by a direct effect on the glands, although the evidence for this is not yet firm. In addition, JH effects, at least in the last larval instar, change from a primarily static response to a tropic response on the brain–prothoracic gland axis at a time shortly before the onset of wandering (Hiruma et al., 1978; Cymborowski and Stolarz, 1979; Safranek et al., 1980). In this section, JH effects on PTTH secretion from the brain and ecdysone secretion from the prothoracic glands will be discussed. The effect of ecdysteroids on CA secretory activity is detailed elsewhere (see Chapter 3.7).

3.8.6.1. Effects of JH on PTTH Secretion

Injection or topical application of JH or its analogs methoprene (effective in Bombyx) or hydroprene (effective in Manduca), at an appropriate period during the feeding stage of the last larval instar of Lepidoptera, delays the onset of the prepupal period in Manduca (Nijhout and Williams, 1974b; Safranek et al., 1980; Rountree and Bollenbacher, 1986; Watson and Bollenbacher, 1988), Spodoptera littoralis (Cymborowski and Stolarz, 1979), and Bombyx (Akai et al., 1973; Sakurai and Imokawa, 1988; Gu et al., 1997). In Manduca, the prothoracic glands from hydroprene- or JH I-treated larvae are capable of responding to PTTH in vitro. Therefore, the delay is supposed to be due to the inhibition of PTTH release from the brains (Rountree and Bollenbacher, 1986). In Bombyx, JH analog (JHA) application before day 3 of the fifth instar results in a delay of the onset of wandering, while its application on day 3 produces dauer (permanent) larvae that never undergo metamorphosis, but die after surviving for more than 2 weeks (Akai et al., 1973; Sakurai and Imokawa, 1988). In such larvae, the prothoracic glands retain their sensitivity to PTTH, when assessed by an in vitro PTTH challenge. In addition, PTTH injection 10 days after the JHA treatment restores spinning behavior and is followed by normal pupation, just as if the days between the JHA treatment and PTTH injection did not exist (Sakurai and Imokawa, 1988). These data indicate that JHA can disturb the program of PTTH secretion in the brain if JHA is applied at a critical time point in the fifth instar. If JH suppresses PTTH release, then what is the effect of JH on the brain–prothoracic gland axis in the penultimate instar when the JH titer is high? Removal of the corpora allata from early penultimate instar larvae results in the induction of precocious pupation. In Mamestra and Bombyx, the hemolymph ecdysteroid titer peaks on day 3 of the fourth instar and induces larval ecdysis. Allatectomy abolishes the appearance of the ecdysteroid peak and the ecdysteroid titer remains at low levels for several days until the precocious onset of wandering (Hiruma, 1986; Ohtaki et al., 1986). The prothoracic glands, after allatectomy, retain their responsiveness to PTTH, indicating that the decline of the JH titer after allatectomy may act on the brain to alter the PTTH secretion program for larval ecdysis. JH application to the allatectomized larvae increases the hemolymph ecdysteroid concentration (Ohtaki et al., 1986), as well as the basal secretory activity of the prothoracic glands within 24 h of the JHA treatment (Sakurai, unpublished data). In

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addition, the brains of early penultimate instar larvae of M. brassicae contain higher PTTH activity than do those of allatectomized larvae (Hiruma, 1986). This may be due to an accumulation of PTTH in the brain, the result of a declining JH titer suppressing PTTH release. These observations suggest that in the penultimate instar, JH may serve to maintain the brain to produce and/or release PTTH, and the brain ceases PTTH release in the absence of JH in the hemolymph. A similar situation is found in the late feeding stage of the last larval instar of Manduca and Mamestra. Allatectomy delays their pupation by delaying the appearance of the ecdysteroid peak in the prepupal period (Hiruma, 1986; Sakurai, 1990). This could be interpreted as a result of a delay in PTTH release due to low JH titer, although there are no data documenting either the release of PTTH from the brain, nor the PTTH titer in the hemolymph. Only one instance of the direct measurement of PTTH in the hemolymph is available to assess these effects of JH (Mizoguchi, 2001). On days 0–2 of the last larval instar in Bombyx, topically applied JH I increases the PTTH concentration in hemolymph significantly. When day 0 larvae were treated with 3 mg JH I, the PTTH titer increased from 28 to 50 pg ml1 in 12 h. Whether or not this increase is of significant biological meaning, it is at least clear that JH does not inhibit PTTH secretion, even in early fifth instar. This is contrary to the data of Rountree and Bollenbacher (1986) who showed that JH inhibits PTTH release in feeding last instar larvae in Manduca. Rather, JH seems to act as a positive regulator of brain PTTH secretion, as has been proposed in Galleria mellonella (Sehnal et al., 1981). Then, how can we interpret the earlier observations that JH induces dauer larvae or causes a delay in the onset of wandering if applied early in the fifth instar? Induction of dauer larvae require an injection of 10 mg JH I (Akai and Kobayashi, 1971) or 1 mg methoprene (Sakurai and Imokawa, 1988). However, the highest titer of JH in the last instar is less than 10 ng JH I equivalent ml1 (Fain and Riddiford, 1975; Baker et al., 1987; Niimi and Sakurai, 1997), which indicates that although the dose of JH is pharmacological, JH nevertheless disturbs the PTTH release program in the brain, directly or indirectly, through an unknown nervous network participating in the control of PTTH cells. Apart from the effects of JH on PTTH release, it is obvious that the brain also responds to an increase and decrease in both ecdysteroid and JH. Quoting Gilbert et al. (2000), ‘‘ecdysteroids and JHs interact and we cannot study one without considering the other.’’ Ecdysteroids and JH must affect the production and

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release of PTTH, although the role of JH on the brain is still obscure due to the lack of in vitro information on PTTH secretion from the brain. 3.8.6.2. Effects of JH on Ecdysteroidogenesis

Effects of JH on ecdysteroidogenesis are somewhat complicated and not yet understood. JH stimulation of prothoracic gland activity was first demonstrated in diapausing pupae of H. cecropia; JH injection broke the diapause (Gilbert and Schneiderman, 1959; Williams, 1959). However, the brain may not be involved in the diapause-terminating effects of JH, since JH effects were observed in pupae whose brains had been removed prior to JH treatment (Gilbert and Schneiderman, 1959). This indicates that the brain of a diapausing pupa is not the primary target of the applied JH. Rather, JH appears to directly stimulate the pupal prothoracic glands. In H. cecropia diapausing pupae (Gilbert and Schneiderman, 1959), JH probably stimulated prothoracic glands to secrete ecdysone so that a positive feedback loop was initiated, operating in a manner similar to that demonstrated for the Manduca pupal prothoracic glands (Sakurai and Williams, 1989). In Manduca pupae, ecdysteroids stimulate prothoracic glands of diapausing pupae, but also those in a very early nondiapausing pupal stage. Although JH stimulation of pupal prothoracic glands has been demonstrated in vivo, JH has not been detected in the pupal stage, at least before degeneration of the prothoracic glands. Thus, JH stimulation of the pupal glands cannot occur in the normal insect life cycle. In the last larval instar of Lepidoptera, JH delays the increase in the hemolymph ecdysteroid titer when topically applied to prewandering larvae. However, its effect on the steroid titer changes from inhibitory to stimulatory shortly before or at the wandering stage (Hiruma et al., 1978; Safranek et al., 1980; Sakurai, 1990). In the prepupal stage, JH has been considered to have a stimulatory effect on prothoracic glands. The hemolymph JH titer begins to increase concomitantly with the onset of wandering and reaches a broad peak at the midpoint of the prepupal period, after which it decreases towards pupation (Baker et al., 1987; Niimi and Sakurai, 1997). Removal of the corpora allata from the last instar larvae during the prewandering stage results in a delay of pupation in Mamestra and Manduca (Hiruma, 1980; Hiruma et al., 1999), indicating that the JH peak may affect ecdysteroidogenesis. Neck ligation of Manduca feeding larvae on the day of wandering suppresses pupation, and JHA (hydroprene) application restores the delay (Safranek et al., 1980). The ligation suppressed the

422 Feedback Regulation of Prothoracic Gland Activity

increase in hemolymph ecdysteroid titer after wandering, while the administration of JHA to those larvae increased these levels to that found in intact larvae. However, in vitro treatment of wandering larval glands with JHA fails to stimulate gland activity (Sakurai, 1990). In the tropical moth Omphisa fuscidentalis, larvae enter diapause at a time after the cessation of feeding, but before gut purge, i.e., probably at the wandering stage. Larval diapause lasts for 9 months, during which the secretory activity of prothoracic glands is detectable (Singtripop et al., 1999, 2000). Nevertheless, the hemolymph ecdysteroid titer remains at low levels (less than 10 ng ml1) in the first half of the diapause period, indicating that there is no feedback activation in spite of the satisfaction of a circumstantial requirement, i.e., low secretory activity of the glands and low ecdysteroid titer driving positive feedback. Topical application of JHA (methoprene) to those larvae breaks the diapause with associated increases in ecdysteroid titer and prothoracic gland secretory activity. This activation by JHA is also seen in larvae whose brains have been removed prior to JHA treatment, indicating that JHA does not stimulate brain PTTH release. Transplantation of the prothoracic glands of JHA-treated larvae to nontreated larvae breaks the diapause of the recipients. Successive transplantation after JHA treatment showed that JHA stimulation for only 1 day was enough to stimulate the prothoracic glands (Singtripop and Sakurai, unpublished data), but gland activity began to increase only 7 days after JHA treatment (Singtripop et al., 2000). What occurred in the glands during the 6 days immediately after JHA stimulation? During the diapause period, the prothoracic glands do not respond to either circulating ecdysteroids or the application of exogenous 20E (Singtripop et al., 2002). Second, JHA may not stimulate the glands directly. Thus, an assumption can be made that the glands are suppressed to respond to feedback activation by ecdysteroids, and JH may release the glands from such suppression. Such glands are capable of responding to ecdysteroids very gradually because the initial ecdysteroid titer is fairly low. This seems to account for the above-mentioned disagreement of in vivo and in vitro results for the effects of JHA on prothoracic glands in the prepupal period in Manduca and Mamestra. JH may act on prothoracic glands as a modulator of feedback activation in the wandering stage and the following prepupal period, and therefore JH application restores the increase of hemolymph ecdysteroid titer in the neck ligated larvae but cannot stimulate the glands in vitro.

In the last larval instar before the onset of wandering, prothoracic glands are suppressed by JH, but there have been no in vitro data, except for experiment using Drosophila ring glands (Richard and Gilbert, 1991). Both juvenile hormone bisepoxide (JHB3), a Drosophila JH, and JH III well inhibit the ring glands, but the effective concentration is in the mM range, a rather pharmacological concentration. Application of hydroprene, an effective JH analog in Manduca, to larvae before the onset of wandering inhibits the increase in the hemolymph ecdysteroids that normally occurs with wandering, and the prothoracic gland activity as well (Rountree and Bollenbacher, 1986; Sakurai, 1990). In Bombyx feeding larvae, JHA application on day 3 of the fifth instar induces dauer larvae although the prothoracic glands after JHA treatment exhibit a low, but significant secretory activity that remains at low levels for as long as the larvae survive. By contrast, the gland activity in larvae whose brains have been removed increases autonomously after the operation to a degree sufficient to induce pupation (Sakurai and Imokawa, 1988). This indicates that JHA applied on day 3 might suppress the glands ability to respond to feedback activation, in addition to disturbing the PTTH secretion program of the brain discussed below (see Section 3.8.6.3). Thus, the switchover of the JH effects on prothoracic glands, occurring around the time of wandering, does not mean the switching from direct suppression to activation by JH. Rather, it may be caused by a change in the gland’s responsiveness to feedback regulation. After wandering, JH supports feedback activation and/or interferes with feedback inhibition, although JH may directly suppress the glands before wandering, if the ecdysteroid titer is low. There is no information at the molecular level on the effects of JH on prothoracic glands although JH modulates the 20E-inducible EcR and USP gene expressions in the Manduca epidermis (Hiruma et al., 1999). In day 2 fourth instar larval epidermis, USP-1 is downregulated by 20E, while USP-2 is upregulated. Coexistence of JH I with 20E suppresses the downregulation of USP-1 expression, indicating that JH may act antagonistically against 20E. USP is suggested to bind JH and act as a JH nuclear receptor (Jones and Sharp, 1997; Jones et al., 2001) (see Chapter 3.7), although the binding affinity of JH to USP is very low, and therefore USP is not yet accepted as a true JH nuclear receptor. JH is reported to activate EcR-dependent transcription in Chinese hamster ovary cells (Henrich et al., 2003). 20E activated the reporter gene expression in Chinese hamster ovary cells transfected with plasmids carrying a Drosophila EcR gene, a USP

Feedback Regulation of Prothoracic Gland Activity

gene, and an ecdysteroid-inducible reporter gene with five ecdysone response elements. In this system, the additional presence of JH III enhanced the response. The effective concentration of JH III was 40 mM (10 600 ng ml1), which is much higher than the JH concentration in larval hemolymph so far reported as less than 10 ng ml1, and thus the effect of JH III is only pharmacological. Accordingly, it is not known for sure if JH interacts directly with the EcR/USP heterodimer in situ to modulate the function of 20E-EcR/USP, but those reports indicate that JH is at least capable of binding to EcR/USP. If this is the case in the prothoracic glands, then JH may modulate the action of 20E by binding to USP-2 and reducing the feedback inhibition by 20E-EcR/USP-2, at least, in the prepupal period when the 20E titer is high enough to induce the complementary expressions of USP-1 and USP-2. If so, the delay in pupation caused by removal of corpora allata at the wandering stage could be interpreted as implying that JH in the prepupal period suppresses the feedback inhibition by suppressing the USP-2 expression by 20E, so that the glands secrete ecdysteroids until the peak titer. In the absence of JH, ecdysteroidogenesis is suppressed by 20E, resulting in an ecdysteroid titer that does not increase. The result is a delay in pupation. This interpretation of JH interaction with USP appears to account well for the fact that the USP-2 upregulation in Manduca prothoracic glands occurs only 1 day before the peak in ecdysteroid titer. In the prepupal period, the threshold concentration of 20E needed to induce USP-2 in the presence of JH may be higher than that in the absence of JH, and it is therefore of interest to examine the effects of JH on USP gene expression in the prothoracic glands. 3.8.6.3. Growth-Blocking Peptide–Corpora Allata–Prothoracic Gland Pathway

Endocrine control of larval growth could be more complicated than previously considered. There is another factor, besides PTTH, 20E and JH, that is indirectly involved in the control of ecdysteroidogenesis. The factor is growth-blocking peptide (GBP), an insect cytokine, which was first identified from the hemolymph of lepidopteran larvae that had been parasitized by a wasp (Hayakawa, 1991). When last (sixth) instar larvae of the armyworm Pseudaletia separata were parasitized by the parasitoid wasp Cotesia kariyai, larval growth was delayed so that the parasitoid wasp larvae had sufficient time to complete larval growth inside the host larvae and escape from them in order to pupate. The GBP concentration in the hemolymph is high in the penultimate instar and gradually declines early in

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the last larval instar. During normal growth, JH esterase in the hemolymph increases in early last instar larvae (Sparks et al., 1983; Baker et al., 1987), and the GBP concentration decreases concomitantly with the increase in esterase activity (Ohnishi et al., 1995). GBP is produced by the larval fat body and acts to suppress JH esterase gene expression. Accordingly, suppression of GBP expression results in increased esterase activity, leading to a decline in JH concentration in the hemolymph. In the parasitized larvae, the GBP gene is activated by parasitization, and the elevated GBP suppresses the decline of JH titer by repressing the JH esterase gene (Hayakawa, 1990), which causes the hemolymph JH titer to remain high and thereby suppress the reactivation of the prothoracic glands. Thus, the parasite delays the growth of the host larvae. GBP also acts on epidermal cells by upregulating the DDC gene, resulting in an enhanced production of dopamine (Noguchi and Hayakawa, 1996; Noguchi et al., 2003). Dopamine is released into the hemolymph and acts to enhance JH secretion from the corpora allata in early last instar larvae (Granger et al., 1996) (see Chapter 3.7). These all figure in the sequential control of prothoracic gland activity (Figure 1). In the early last larval instar, the GBP gene is downregulated, although the mechanism is not known, resulting in a decrease in hemolymph GBP concentration. This decrease causes a downregulation of the DDC gene and results in a low dopamine concentration in the hemolymph. Although dopamine may not be the sole factor to maintain activity in the corpora allata, nevertheless, this activity may decline due to a low dopamine concentration. Simultaneously, JH esterase activity increases and as a result, the JH titer declines sharply, although it remains to be seen whether dopamine acts directly to suppress esterase gene expression or if the decreasing GBP titer directly elicits the upregulation of the JH esterase gene in the fat body. In addition to the GBP pathway, 20E is known to maintain or even stimulate the JH synthetic activity of the corpora allata (Whisenton et al., 1987a, 1987b; Gu and Chow, 1996). Accordingly, the extremely low ecdysteroid titer in the early fifth instar (Bollenbacher et al., 1981; Sakurai et al., 1998) also contributes to the decrease in activity in the corpora allata. Finally, the declining JH titer releases the prothoracic glands from their suppression by JH and allows them to begin ecdysteroidogenesis, as well as to respond to PTTH (Sakurai et al., 1989a). Because GBP is tightly involved in the changes in JH titer, the GBP–JH–prothoracic gland axis could be of importance in the control of prothoracic gland activity and therefore in

424 Feedback Regulation of Prothoracic Gland Activity

the hormonal control of insect growth and development. Since the parasitization of P. separata by the parasitoid A. kariyai appears to affect the host brain in a manner to suppress the PTTH release (Tanaka et al., 1987), GBP could be affecting the brain PTTH cells. There are conditions other than parasitization that delay larval growth, including low temperature (Mala´ et al., 1987; Ohnishi et al., 1995), insufficient food supply (Bounhiol, 1938; Nijhout and Williams, 1974a; Morita and Tojo, 1985; Miner et al., 2000), and injury (O’Farrell et al., 1960; Kunkel, 1977; Mala´ et al., 1987). When last instar P. separata larvae are transferred from 25 to 10  C, pupation is delayed by 11 days, and after the transfer, the GBP concentration is elevated and maintained at high levels for a certain period (Ohnishi et al., 1995), indicating that the cold-stress suppressed, or even upregulated, the GBP gene in the fat body, which probably led to the delayed activation of prothoracic glands through a suppression of the decline of JH titer. Chilling of last instar G. mellonella larvae elicits an extra larval molt or a delay of pupation (Mala´ et al., 1987). It is of interest that GBP is involved in the delay of larval growth under certain stressed circumstances. In the starved larvae, the feedback regulation of prothoracic gland activity seems to be more complicated. Starvation in last instar larvae of Manduca results in the maintainance of a high level of hemolymph JH, rather than the decline observed in normal feeding larvae (Cymborowski et al., 1982), apparently due to the suppression of the decline in hemolymph JH esterase activity. Thus, the resulting high JH level may modulate the brain– prothoracic gland axis as mentioned above. This could be accounted for by the suppression of GBP expression that normally ceases after the last larval molt. However, suppression of larval growth by starvation is not simply caused by the suppression of prothoracic gland activity. Starvation suppresses the release of growth factors, like bombyxin and IDGFs, which are needed for normal growth. Nevertheless, 20E is required simultaneously to sustain larval growth (Nijhout and Grunert, 2002). Accordingly, the horizontal regulation of prothoracic glands appears to participate in helping the animal survive unexpectedly severe circumstances. For more than four decades, injury has been recognized to delay larval growth, known as the injury effect. Following an operation such as the sham operation for allatectomy or autotomy of larval legs, the larval or even pupal period is usually prolonged by one or more days over that of intact animals. Similar sham operations on the last instar larvae of G. mellonella either prolongs the intermolt

periods or produces supernumerary larvae. The injury is supposed to stimulate JH production (Mala´ et al., 1987). In the sixth instar larvae of the cockroach Blatta orientalis, autotomy results in a delay of ecdysis by 4 days and also delays the appearance of the hemolymph ecdysteroid peak by 4 days (Kunkel, 1977), indicating that injury alters the ecdysone secretion program by affecting the brain– prothoracic gland axis. These two results are well explained by the operation of the GBP pathway (Figure 1). Injury elicits an increase in GBP titer, which stimulates the corpora allata to produce JH and also suppresses JH esterase expression, resulting in a high JH titer. The high titer suppresses the brain–prothoracic gland axis, delaying the appearance of the peak ecdysteroid titer. As a result, injury either delays ecdysis or induces supernumerary larvae. Thus, it seems highly possible that the humoral factor induced by injury is GBP, and that the GBP system is the cause for the delay in development.

3.8.7. Role of Feedback Regulation in Larval Growth and Metamorphosis: Meaning of the Shapes of the Ecdysteroid Peak In contrast to the pupal (pharate adult) stage, during which the prothoracic glands degenerate and are unable to produce additional ecdysone, at the end of every larval stage including the last larval instar, gland activity decreases to a very low level. However, the degree of inactivation achieved is different at the end of each of at least the third, fourth, and fifth (last) instars of Manduca (Bollenbacher et al., 1987; Smith and Pasquarello, 1989; Rybczynski and Gilbert, 1994; Smith, 1995) and Bombyx (Okuda et al., 1985; Sakurai et al., 1989b; Gu et al., 1996, 2000; Takaki and Sakurai, 2003). Therefore, the mode of feedback inhibition of prothoracic glands may change developmentally. When the ecdysteroid titer peaks occurring at the end of the third, fourth, and fifth larval instars and pupal stage are compared to each other, it is easily recognized that the peak titer, as well as its width, are developmental stage specific in Manduca (Bollenbacher et al., 1981) and Bombyx (Kiguchi and Agui, 1981; Mizoguchi et al., 2001; Takaki and Sakurai, 2003). In Bombyx, the peak width at the point of 50% peak height in the third, fourth, and fifth instars and pupal stage are 7 h (peak value of 1.2 mM), 9 h (1 mM), 18 h (3.2 mM), and 24 h (15.4 mM), respectively. The duration of the period with the titer above 1  107 M, a value similar to the binding affinity of 20E for the EcR/USP heterodimer, is 18 h, 32 h, 3 days, and 6

Feedback Regulation of Prothoracic Gland Activity

days, respectively (estimated from Takaki and Sakurai (2003) for the third and fourth instars, and from Mizoguchi et al. (2001) for the fifth instar and pupal periods). Thus, the shape of the ecdysteroid titer peak prior to the stationary molt in the third and fourth instars is sharper and narrower than those in the prepupal and pupal period. At the pupal molt the peak value is threefold higher than that for the stationary molt. This value in the pupal period is fivefold higher than that in the prepupal period. This is also the case in Manduca (Bollenbacher et al., 1981, 1987; Warren and Gilbert, 1986). Although little attention has been focused on such differences so far, these changes must possess an important meaning in the control of insect growth and metamorphosis. In Bombyx, the composition of hemolymph ecdysteroids is strikingly different at the peak titers occurring between the fourth instar and prepupal period (Kamimura and Kiuchi, 2002). At day 2.5, 0.5 days prior to the peak in hemolymph ecdysteroids occurring in the fourth instar that lasts 4.5 days, the 20E concentration is fivefold higher than that of ecdysone, while at the peak, the 20E titer is 17 times as great as that of ecdysone. In contrast, 0.5 days prior to the peak titer in the prepupal period, ecdysone is the predominant ecdysteroid, as the 20E concentration is approximately 70% that of ecdysone. 20E becomes predominant by the time of the peak titer, although 20E concentrations are only 30% higher than ecdysone. In addition, the absolute concentrations of 20E at peak titer are 620 ng ml1 (1.3 mM) in the fourth instar and 480 ng ml1 (1 mM) in the fifth instar. In Manduca prothoracic glands of day 5 fifth instar larvae, 1 mM 20E significantly suppresses USP-1 while at the same time it stimulates USP-2 expression (Song and Gilbert, 1998). The fourth instar glands are also inhibited by 20E in vitro, similar to those of the fifth instar (Sakurai and Gilbert, unpublished data). These composite data from Manduca and Bombyx indicate that in the fourth instar, 20-monooxygenase activity sharply increases when prothoracic gland activity increases, and therefore secreted ecdysone is promptly converted to 20E, which provides for an acute induction of USP-2 and thereby elicits a strong negative feedback along with the increased sensitivity of the glands to 20E (Takaki and Sakurai, 2003). By contrast, in the prepupal period, 20-monooxygenase activity may only gradually increase, while the degree of feedback inhibition is not so strong as compared with that in the fourth instar. While receiving a weak feedback inhibition by 20E in the early period of feedback inhibition, the prothoracic glands are capable of responding to PTTH in the presence of 20E (Sakurai

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and Williams, 1989; Song and Gilbert, 1998), and therefore the glands may be able to respond to PTTH and secrete ecdysone even in the face of the feedback inhibition that occurs before the peak titer in the prepupal period. The degree of sharpness of both the increase and decrease in the titer cannot be accounted for only by feedforward (PTTH) and feedback factors. As discussed elsewhere in this volume (see Chapters 3.2, 3.3, and 3.7) and above, various factors are involved in the control of prothoracic gland secretory activity resulting in both upregulation and downregulation (Figure 1). JH acts as a modulator in a stage-specific manner. Nerves from the prothoracic ganglion suppress gland activity in at least some insects (Okajima and Kumagai, 1989; Richter and Bo¨ hm, 1997). Decreases in the titer also depend on ecdysteroid metabolism, starting with the conversion of ecdysone to 20E by the 20-monooxygenase followed by 26-hydroxylation mediated by the 26-hydroxylase that is induced by 20E (Hoffman et al., 1974; Williams et al., 1997). Thus, the stage-specific expression of these enzymes is also tightly involved in the decrease in the ecdysteroid titer. The peak value, composition of ecdysteroids, and duration of ecdysteroid titer above a certain concentration are controlled precisely in a stage-specific manner, but the overall output of the gland is dominant and governs every aspect, directly or indirectly, of insect postembryonic development. Ecdysone has been regarded as an inactive prohormone because its affinity to EcR/USP is far less than that of 20E (Minakuchi et al., 2003). This low affinity well accounts for its very low biological activity measured using an in vitro system (Cherbas et al., 1980) where exogenous 20-monooxygenase activity is not a factor. A hormonal role for ecdysone is still conjectural, but one has been suggested over the last three decades (Mandaron, 1973; Tanaka and Takeda, 1993; Tanaka and Naya, 1995; Gilbert et al., 1996; Kamimura and Kiuchi, 2002). Nevertheless, 20E must be the main feedback factor in the brain–prothoracic gland axis.

3.8.8. Future Directions In the study of feedback regulation of the endocrine system, one of the advantages of insects over mammals is that the regulation of most endocrine systems is tightly associated with embryonic and postembryonic development, including growth, pupal– adult differentiation, and cell death. In vertebrates, although hormonal regulation is seen in embryonic development, feedback and feedforward regulation

426 Feedback Regulation of Prothoracic Gland Activity

in the endocrine system is focused to a great extent on the adult stage and therefore the regulation is rather stereotypic. In insects, all the cells of the body are targets of two hormones, 20E and JH, regardless of their developmental stages. The regulation of prothoracic gland ecdysteroidogenesis is believed to be under the control of a complicated regulatory network. The regulatory mechanism of prothoracic gland activity includes almost every aspect found in a mammalian system, i.e., involving tropic and static factors, as well as a horizontal network of the control, neural control, and feedback regulation. At the cellular level, those factors may act through signal transducing pathways mediated by second messengers, a variety of kinases, and different combinations of transcription factors, including the nuclear receptors, EcRs and USPs, and their coactivators. All these factors could be involved in the regulation of gene expression, protein synthesis, and the transport of steroid intermediates in the prothoracic gland cells. As a result of the integration of all of those regulatory pathways, ecdysone secretion is either stimulated or suppressed. With the recent advances of analytical equipment and techniques, macrolepidopteran prothoracic glands (Manduca) are sufficiently large in mass to perform biochemical and pharmacological analysis of the aforementioned complicated regulation. However, Drosophila is different from other insects; for example, the involvement of JH in the stationary molt is not understood. Nevertheless, precocious metamorphosis can be induced by genetic manipulation (Bialecki et al., 2002), indicating that there should be a control mechanism to ensure the stationary molt. The most sophisticated genetic manipulation is available in Drosophila, which will be of great help in understanding the complicated feedback regulation systems in the endocrine systems of insects.

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ecdysteroid biosynthesis in the prothoracic glands of Manduca sexta. Insect Biochem. Mol. Biol. 25, 591–602. Song, Q., Gilbert, L.I., 1997. Molecular cloning, developmental expression, and phosphorylation of ribosomal protein S6 in the endocrine gland responsible for insect molting. J. Biol. Chem. 272, 4429–4435. Song, Q., Gilbert, L.I., 1998. Alterations in ultraspiracle (USP) content and phosphorylation state accompany feedback regulation of ecdysone synthesis in the insect prothoracic gland. Insect Biochem. Mol. Biol. 28, 849–860. Sparks, T.C., Hammock, B.D., Riddiford, L.M., 1983. The haemolymph juvenile hormone esterase of Manduca sexta (L.): inhibition and regulation. Insect Biochem. 13, 529–541. Steel, C.G.H., 1975. A neuroendocrine feedback mechanism in the insect moulting cycle. Nature 253, 267–269. Steel, C.G.H., Harmsen, R., 1971. Dynamics of the neurosecretory system in the brain of an insect, Rhodnius prolixus, during growth and molting. Gen. Comp. Endocrinol. 17, 125–141. Suzuki, C., Ishizaki, H., 1986. Prothoracicotropic hormone bioassay: Bombyx larval assay. Int. J. Invertebr. Reprod. Devel. 10, 259–274. Takaki, K., Sakurai, S., 2003. Regulation of prothoracic gland steroidogenic activity leading to pupal metamorphosis. Insect Biochem. Mol. Biol. 33, 1189–1199. Tanaka, T., Agui, N., Hiruma, K., 1987. The parasitoid Apanteles kariyai inhibits pupation of its host, Pseudaletia separata, via disruption of prothoracicotropic hormone release. Gen. Comp. Endocrinol. 67, 364–374. Tanaka, Y., Naya, S., 1995. Dietary effect of ecdysone and 20-hydroxyecdysone on larval development of two lepidopteran species. Appl. Entomol. Zool. 30, 285–294. Tanaka, Y., Takeda, S., 1993. Ultranumerary larval ecdysis of the silkworm, Bombyx mori induced by ecdysone. Naturwissenschaften 80, 131–132. Truman, J.W., 1972. Physiology of insect rhythms. 1. Circadian organization of the endocrine events underlying the moulting cycle of larval tobacco hornworms. J. Exp. Biol. 57, 805–820. Wang, S.F., Li, C., Zhu, J., Miura, K., Miksicek, R.J., et al., 2000. Differential expression and regulation by 20hydroxyecdysone of mosquito ultraspiracle isoforms. Devel. Biol. 218, 99–113. Warren, J.T., Gilbert, L.I., 1986. Ecdysone metabolism and distribution during the pupal–adult development of Manduca sexta. Insect Biochem. 6, 65–82. Warren, J.T., Gilbert, L.I., 1996. Metabolism in vitro of cholesterol and 25-hydroxycholesterol by the larval prothoracic glands of Manduca sexta. Insect Biochem. Mol. Biol. 26, 917–929. Warren, J.T., Petryk, A., Marque´ s, G., Jarcho, M., Parvy, J.-P., et al., 2002. Molecular and biochemical characterization of two P450 enzymes in the ecdysteroidogenic

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3.9 Hormonal Control of Reproductive Processes A S Raikhel, University of California, Riverside, CA, USA M R Brown, University of Georgia, Athens, GA, USA X Belles, Institut de Biologia Molecular de Barcelona (CSIC), Spain ß 2005, Elsevier BV. All Rights Reserved.

3.9.1. Introduction 3.9.2. Endocrine Control of Female Reproduction 3.9.2.1. Evolution of Endocrine Control of Vitellogenesis and Egg Development 3.9.2.2. Yolk Protein Precursors 3.9.2.3. Mechanisms of Juvenile Hormone Action in Vitellogenesis 3.9.2.4. Molecular Mechanisms of Ecdysteroid Action in Vitellogenesis 3.9.2.5. Molecular Endocrinology of Vitellogenesis in the Cyclorrhaphan Diptera 3.9.2.6. Hormones and Ovarian Maturation 3.9.2.7. Hormone Action in Female Accessory Glands 3.9.2.8. Oviposition 3.9.2.9. Peptide Hormones Involved in Female Reproduction 3.9.3. Hormones and Male Reproduction 3.9.3.1. Spermatogenesis 3.9.3.2. Male Accessory Gland Function 3.9.4. Future Directions

3.9.1. Introduction The years following the 1985 publication of Comprehensive Insect Physiology, Biochemistry and Pharmacology have been marked by stunning developments in insect science. A technological revolution in biochemistry, molecular biology, and genetics has swept all areas of biological science, and has profoundly influenced insect science as well. With this technological revolution, the small size of insects is no longer a barrier to discovering their biochemical make-up, cloning or characterizing genes, or uncovering genetic and hormonal signals governing their functions. Sequencing of insect genomes, particularly that of Drosophila melanogaster, has led to the identification of previously unknown genes and to the development of functional genomic approaches that lead to further elucidation of genetic regulatory networks. During this period, insect endocrinology has shifted its focus from physiology and biochemistry to molecular biology and genetics. The latter are the subjects of the present volume. This chapter specifically reviews the progress that has been achieved since 1985 in our understanding of the hormonal control of insect reproduction. Particular attention is paid to those areas where significant progress has been made at the molecular and genetic levels.

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There are a number of recent reviews on hormonal regulation of yolk protein (yp) genes (Raikhel et al., 2003; Belles, 2004; Bownes, 2004; Wang et al., 2004); vitellogenins and their processing (Sappington et al., 2002; Telfer, 2002; Tufail et al., 2004); nonvitellogenin yolk proteins (Bownes and Pathirana, 2002; Telfer, 2002; Masuda et al., 2004; Yamahama et al., 2004), and the cell biology of the insect fat body (Giorgi et al., 2004).

3.9.2. Endocrine Control of Female Reproduction 3.9.2.1. Evolution of Endocrine Control of Vitellogenesis and Egg Development

The insect female reproductive system consists of the ovaries, which contain ovarioles, oviducts, spermatheca, accessory glands, vagina, and ovipositor (Davey, 1985; Chapman, 1998). Ovarioles are the egg producing units in the ovary. Typically, the insect ovary has four to ten ovarioles; however, some species contain many more ovarioles. Ovarioles are tubular and consist of both somatic and germline cells. At the apex of each ovariole, a germarium houses the primary germline cells. The follicles or egg chambers form within the germarium

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and continue to mature along the ovariole tube. There are three types of ovaries in insects (Bu¨ning, 1994). Ovaries of primitive groups of insects contain panoistic ovarioles with egg chambers consisting of the oocytes surrounded by the follicular epithelium (panoistic ovarioles). In more advanced insects, where the structure of ovarioles is more complex (meroistic), some germ cells are set aside to form nurse cells that produce massive amounts of numerous products for the developing oocyte. In polytrophic meroistic ovarioles, each egg chamber contains its own group of nurse cells connected to the oocyte. Follicle cells surround the oocyte and nurse cells. In telotrophic meroistic ovaries, each ovariole contains the trophic chamber with large groups of nurse cells. The trophic chamber is connected to the egg chambers by nutritive cords. Egg chambers of telotrophic ovarioles contain only the oocyte surrounded by the follicle cells. Endocrine control of female reproduction is governed by different types of hormones: neuropeptides (see Chapter 3.10), juvenile hormones (JHs; see Chapter 3.7), and ecdysteroids (see Chapters 3.3 and 3.5). In keeping with common practice among researchers in the field, ecdysteroid is used as the generic term for steroidal insect molting hormones, reserving the term ecdysone for the specific chemical compound 2b, 3b, 14a, 22R, 25-pentahydroxy-5b-cholest-7-en-6-one, originally known as a-ecdysone. The abbreviation 20E will be used to refer to 20-hydroxyecdysone (2b, 3b, 14a, 20R, 22R, 25-pentahydroxy-5b-cholest-7-en-6-one), the ecdysone metabolite believed to serve as the active hormone in most well-characterized responses. Individually, or in concert, the regulation of particular events during female reproduction by these hormones has evolved along with their effects on molting and metamorphosis. Control of female reproduction in Apterygota remains the most enigmatic. Studies of the firebrat, Thermobia domestica (Zygentoma), have shed some light on the hormonal regulation of vitellogenesis and ovarian development in primitive apterous insects (Bitsch and Bitsch, 1984; Bitsch et al., 1986). In this insect, molting occurs continually into the adult stage, and oogenesis is coordinated with the ecdysteroid regulated adult molting cycle (Bitsch et al., 1986). Treatment with the anti-allatal drug precocene blocks ovarian maturation, which indicates its dependence upon juvenile hormone secreted by the corpora allata (CA) (Bitsch and Bitsch, 1984; Bitsch et al., 1986). Further studies are required in order to understand the precise roles of JH and ecdysteroids in regulating vitellogenesis and oogenesis in apterous insects.

It is well established that major events of reproduction in all insect orders with incomplete metamorphosis (Hemimetabola) are governed by JH. In the orders with complete metamorphosis (Holometabola), control strategies have evolved differently. In beetles (Coleoptera), JH remains the major regulatory hormone of reproductive events. In Hymenoptera, the role of JH is elaborated in eusocial species having a single or a few reproductive females in a colony (see Chapter 3.13). In Lepidoptera, female reproduction is regulated either by JH or ecdysteroids. In many such species, egg maturation occurs during the pharate adult stage and requires coordination with hormonal signals controlling metamorphosis. In dipteran insects, mosquitos, and flies, ecdysteroids have the leading role as hormonal regulators, but JH has an important role as a regulator in dipteran females in preparing reproductive tissues for ecdysteroid mediated events, such as vitellogenesis. For all insects, neuropeptides play a key role regulating the production of JH and ecdysteroids. A summary of the primary hormones involved in reproduction according to the phylogeny of insect orders is presented in Figure 1; for some orders, this is not known.

Figure 1 Utilization of juvenile hormone or ecdysteroids as major regulators of vitellogenesis and reproduction among insect orders. Some hemipterans show incomplete dependence on JH. In hymenopterans, JH plays a vitellogenic role in nonsocial and primitive social groups, but not in advanced groups (see Chapter 3.13). In lepidopterans, those species that begin vitellogenesis after adult emergence are JH dependent, and those in which vitellogenesis proceeds between pupal and adult stages are partially dependent on JH, whereas species in which vitellogenesis proceeds within or before the pupal stage are independent of JH. Dipterans, in general, are ecdysteroid dependent, although JH may play an accessory role in vitellogenesis (Modified from Belles, X., 2004. Vitellogenesis directed by juvenile hormone. In: Raikhel, A.S., Sappington, T.W. (Eds.), Reproductive Biology of Invertebrates, Vol. 12, Part B: Progress in Vitellogenesis. Science Publishers, Enfield, USA/Plymouth, UK, pp. 157–198).

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3.9.2.1.1. Juvenile hormone directed female reproduction In the adult females of Hemimetabola (Dictyoptera to Hemiptera) and Coleoptera, JH is the main regulator and pleiotropically controls most aspects of female reproduction (Figure 2). The major role of JH in reproduction is to regulate vitellogenin (Vg) gene expression in the fat body, generally, and in ovarian follicular epithelium (Wyatt and Davey, 1996; Engelmann, 1983, 2003; Belles, 2004). Some gryllid and hemipteran species

Figure 2 The pleiotropic role of juvenile hormone in insect reproduction. ER, endoplasmic reticulum; Vg, vitellogenin. (Reproduced with permission from Engelmann, F., 2003. Juvenile hormone action in insect reproduction. In: Henry, H.L., Norman, A.W. (Eds.), Encyclopedia of Hormones, Vol. 2. Academic Press, San Diego, pp. 536–539.)

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show an incomplete dependence upon JH (Wyatt and Davey, 1996; Strambi et al., 1997). Cockroaches (Dictyoptera) are the classical model for studies of JH dependent vitellogenesis. In the oviparous Periplaneta americana (Blattidae), Weaver and Edwards (1990) have shown that allatectomy or treatment with inhibitors of JH synthesis blocks Vg production, oocyte growth, and ootheca formation, whereas JH treatment restores these processes. The regulation of vitellogenesis by JH has been most thoroughly studied in Blattella germanica (Blattellidae). In this typical JH dependent species, in which the females carry the ootheca externally until the first inside larvae emerge, JH production (Belles et al., 1987), Vg titers (Martı´n et al., 1995), and Vg mRNA levels (Comas et al., 1999) show cyclical and approximately parallel patterns during the reproductive cycle (Figure 3). The pseudoviviparous cockroach, Leucophaea maderae, has been one of the favorite models for biochemical studies of JH regulation of reproduction. Effective doses to induce vitellogenesis in females in vivo range from 1 mg of JH I to 25 mg of JH III in vivo. Methoprene, a potent JH analog (JHA), induces vitellogenesis in adult males, as well (Don-Wheeler and Engelmann, 1997). In another pseudoviviparous cockroach, Blaberus discoidalis, allatectomy prevents ovarian maturation that can be restored by JH III treatment (Keeley and McKercher, 1985). In decapitated adult females, methoprene or JH III induces Vg protein synthesis (Keeley et al., 1988) and a 6.5 kb mRNA, presumably corresponding to the Vg transcript (Bradfield et al., 1990). The neuropeptide, hypertrehalosemic hormone, enhances the vitellogenic

Figure 3 Production rates of juvenile hormone (JH) from the corpora allata, vitellogenin (Vg) from the periovarial fat body, and fat body Vg mRNA during the first reproductive cycle in the cockroach Blattella germanica. (Based on data from Belles, X., Casas, J., Messenguer, A., Piulachs, M.D., 1987. In vitro biosynthesis of JH III by the corpora allata of Blattella germanica (L.) adult females. Insect Biochem. 17, 1007–1010; Martin, D., Piulachs, M.D., Belles, X., 1995. Patterns of hemolymph vitellogenin and ovarian vitellin in the German cockroach, and the role of juvenile hormone. Physiol. Entomol. 20, 59–65; and Martin, D., Comas, D., Piulachs, M.D., Belles, X., 1998. Isolation and sequence of a partial vitellogenin cDNA from the cockroach Blattella germanica (L.) (Dictyoptera, Blattellidae), and characterization of the vitellogenin gene expression. Arch. Insect Biochem. Physiol. 38, 137–146.)

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effects of JH (Lee and Keeley, 1994a). In Nauphoeta cinerea, in which hemolymph Vg levels show a cyclic pattern during the gonadotropic cycle, JH I, II, and III can induce Vg production (Buschor and Lanzrein, 1983). In Dermaptera, the CA are required for reproduction in female earwigs, Anisolabis maritima, as first shown by Ozeki (1951; see Rankin et al., 1997), and JH is the main gonadotropic hormone in female Euborellia annulipes, which have vitellogenic cycles (Rankin et al., 1997). Rouland et al. (1981) reported that allatectomized females Labidura riparia do not produce vitellogenic proteins, and 0.1 mg of farnesol, a precursor of JH, restores vitellogenesis. In Orthoptera, the regulation of vitellogenesis by JH has been demonstrated in a number of locusts and grasshoppers (Acrididae) through classical experiments involving allatectomy and JH treatment (Wyatt, 1991; Wyatt and Davey, 1996). The most thoroughly studied orthopteran species has been Locusta migratoria, in which Vg synthesis is cyclic (Chinzei and Wyatt, 1985). Induction of Vg synthesis in female L. migratoria by JH homologs is only achieved with repeated doses or with high doses coinjected with a JH esterase inhibitor (Wyatt et al., 1987), whereas synthetic JHAs at doses between 2 and 30 mg have a much more potent vitellogenic action (Edwards et al., 1993; Zhang et al., 1993). Neuropeptides may also be involved, since a brain factor enhances vitellogenesis induced by JH in fat body incubated in vitro (Glinka et al., 1995). JH titer, and Vg protein and mRNA levels have been described during the reproductive cycle of the locust Schistocerca gregaria (Mahamat et al., 1997) and the grasshopper Romalea microptera (¼ Romalea guttata) (Borst et al., 2000). In contrast to other orthopteroids, the phasmid Carausius morosus is independent of JH for vitellogenesis (Bradley et al., 1995). Allatectomy does not prevent production of viable eggs in this insect, as first reported by Pflugfelder in 1930, and later confirmed with antibodies against C. morosus Vg that detected its production in allatectomized and intact females (Bradley et al., 1995). However, it is not clear whether the independence from JH for vitellogenesis applies to all Phasmida. In crickets (Gryllidae), both JH and ecdysteroids are involved in the control of vitellogenesis (review: Strambi et al., 1997). In Acheta domesticus, the CA are necessary for oocyte growth, and JH restores vitellogenesis in allatectomized or decapitated females (Renucci et al., 1987; Loher et al., 1992). Conversely, female Teleogryllus commodus emerging from allatectomized larvae can still produce eggs (Loher and Giannakakis,

1990). Although allatectomy of Gryllus bimaculatus reduced the rate of egg production, oocytes in allatectomized females contained a similar amount of yolk protein as oocytes in controls; JH or JHA restored egg production to a variable extent (Hoffmann and Sorge, 1996). Notably, low doses of ecdysteroids stimulated oocyte growth in females of this species (Chudakova et al., 1982; Behrens and Hoffmann, 1983). These results suggest that in crickets in which allatectomy does not completely suppress oocyte growth, ecdysteroids may play a vitellogenic role (Hoffmann and Sorge, 1996). For Hemiptera, the CA were shown to be necessary for vitellogenesis in Rhodnius prolixus by V.B. Wigglesworth in the 1930s, and this observation has since been confirmed for several other hemipteran species (Wyatt and Davey, 1996; Davey, 1997; Belles, 2004). Later studies demonstrated that allatectomy of R. prolixus does not totally abolish Vg synthesis, but JH treatment does restore normal production (Wang and Davey, 1993). Female Oncopeltus fasciatus chemically allatectomized with precocenes produced the Vg precursor, but its conversion to mature Vg, which is incorporated into the oocytes, did not take place, as shown with electrophoresis of native proteins and immunodiffusion (Kelly and Hunt, 1982). A later study of this species found that the synthesis of two female specific Vg components, 200 and 170 kDa, was inhibited in precocene treated specimens and restored by administration of JH (Martinez and Garcera´ , 1987). This discrepancy may be due to differences in the precocene treatment. Only one study has demonstrated a role for JH in a homopteran species. In the black bean aphid, Aphis fabae, ovarian development begins prenatally in this parthenogenic species, in such a manner that oocytes differentiate in the embryonic germaria, and at emergence each ovariole of a first instar aphid already contains one or two developing embryos. Precocene treatment of aphid nymphs inhibited oocyte development in embryos inside the parental ovaries, whereas JH reversed this inhibition (Hardie, 1987). For Coleoptera, JH is the main gonadotropic hormone, but surprisingly little is known about the reproductive endocrinology of this holometabolous order with the greatest number of insect species. A long day regimen for the Colorado potato beetle, Leptinotarsa decemlineata, leads to reproductive activity and a short day induces diapause. JH or JHA (pyriproxyfen) treatment of short-day females induces Vg synthesis, and the JHA also induces Vg production in last instar larvae (de Kort et al., 1997). Similarly, females of Coccinella septempunctata

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reared on suboptimal artificial diets fail to reproduce, but Vg synthesis can be induced by treatment with JHA (Zhai et al., 1984, 1987; Guan, 1989). In the spruce weevil, Pissodes strobi, treatment of previtellogenic females with JH III induces the precocious appearance of Vg transcripts, as determined by the identification of Vg mRNA (Leal et al., 1997). This study also showed that Vg mRNA accumulates slower in beetles feeding on Sitka spruce trees (Picea sitchensis), which are resistant to P. strobi attack, thus suggesting that this plant contains anti-juvenoid compounds. 3.9.2.1.2. From nonsocial sawflies to eusocial ants and bees Within the Hymenoptera, JH appears to play a role in female reproduction among nonsocial and eusocial species and through this action may affect caste determination in the eusocial groups (see Chapter 3.13). In the primitive suborder Symphyta, only Athalia sawflies have received attention. JH applied to male Athalia rosae induced an increase in Vg production, and ovaries implanted into these males incorporated the Vg (Hatakeyama and Oishi, 1990). Later, it was shown that ovaries of Athalia rosae implanted into males of the closely related species, Athalia infumata, incorporated the heterospecific Vg that was induced in the host by JH III treatment (Hatakeyama et al., 1995). Although these studies did not examine vitellogenesis in females, the results suggest a role for JH in females. The regulation of reproduction by JH varies among the suborder Apocrita, which encompasses a range of species from solitary types to highly social groups like ants and honeybees (for reviews of JH action in social Hymenoptera see Robinson and Vargo, 1997; Hartfelder, 2000; Chapter 3.13). In the primitive eusocial paper wasps (Polistidae), Polistes dominulus (¼ Polistes gallicus) and P. metricus, JH acts as a gonadotropin. Queens or dominant females of Polistes show high JH titers associated with growing ovaries, whereas subordinate females have low JH titers. In general, allatectomy of dominant females leads to a reduction in the dominance status. The bumble bee, Bombus terrestris (Apidae), shows a relatively primitive social structure, and JH titers are high in egg laying queens. A regulatory role for JH is substantiated by the positive correlation between ovary development and rate of JH production in queenless workers (Block et al., 2000). The role of JH remains unresolved among the studies of ants and bees with a complex social structure. JH treatment of isolated virgin queens of the fire ant, Solenopsis invicta (Myrmicinae) leads to wing shedding and oviposition, which is prevented by allatectomy. Subsequent JH treatment of

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allatectomized queens induces these phenomena (see Robinson and Vargo, 1997). Conversely, JH titers are low in reproductive female ants in the genus Diacamma (Ponerine; Sommer et al., 1993). In the highly eusocial honeybee, Apis mellifera (Apidae), Vg synthesis in laying queens was only slightly reduced by allatectomy, and JH treatment increased ovary development only weakly (see Robinson and Vargo, 1997). Exogenous JH administered to queen and workers of A. mellifera advances the timing of vitellogenin appearance (Barchuk et al., 2002); together, these results indicate that JH does not play a primary role in honeybee reproduction. 3.9.2.1.3. Butterflies and moths: endocrine compatibility of metamorphosis and vitellogenesis In Lepidoptera, some species begin vitellogenesis after adult emergence, and studies of Papilionoidea (e.g., Pieris brassicae, Nymphalis antiopa, Polygonia caureum, Vanessa cardui, and Danaus plexippus) and Noctuoidea (e.g., Heliothis virescens, Helicoverpa zea, and Pseudaletia unipuncta) show that JH stimulates vitellogenesis (see Cusson et al., 1994a; Ramaswamy et al., 1997). In P. unipuncta, the release rate of JH from CA in vitro is positively correlated with Vg synthesis (Cusson et al., 1994b). In this species and H. virescens, vitellogenesis is abolished in decapitated females, and JH treatment restores it (Cusson et al., 1994a; Ramaswamy et al., 1997). For other groups of Lepidoptera, egg development proceeds between the pupal and pharate adult stages. Studies of species in the Tortricoidea have shown that decapitation of female Choristoneura fumiferana and C. rosaceana reduces egg production, but Vg remains in the hemolymph (Delisle and Cusson, 1999). Treatment of the decapitated females with methoprene restores egg production. Similarly, treatment of female codling moths (Cydia pomonella) with fenoxycarb, a JHA, did not affect protein yolk content, but it did stimulate chorionation (Webb et al., 1999). In this species, ecdysteroids may have a priming effect on vitellogenesis, since treatment with ecdysteroid agonists (tebufenozide and methoxyfenozide) increased levels of circulating Vg but did not affect its incorporation into the oocytes (Sun et al., 2003). These results suggest that vitellogenesis is not completely dependent on JH and that it plays a primary role in Vg uptake by the oocyte and in chorionation in this family. In the Sphingid (Bombycoidea) Manduca sexta, vitellogenesis starts 3–4 days before adult emergence and proceeds in the absence of the pupal CA; thereafter, JH is necessary to complete oocyte growth and chorionation (Satyanarayana et al.,

438 Hormonal Control of Reproductive Processes

1994). Vg was present in prepupae of both sexes at low levels, had disappeared by pupal ecdysis, and reappeared in the late pharate adult females. Methoprene treatment induced Vg synthesis and Vg mRNA in prepupae and freshly molted pupae, and simultaneous administration of ecdysteroids abolished the vitellogenic action of JH (Satyanarayana et al., 1994). The pyraloid moths studied to date have a similar reproductive physiology. In Plodia interpunctella, the declining ecdysteroid titer triggers vitellogenesis in early pupae, and ecdysteroid treatment inhibits vitellogenin uptake by the oocytes (Shirk et al., 1992). Similarly, vitellogenesis in Diatrea grandiosella proceeds in the absence of ecdysteroids, whereas choriogenesis is completed with JH in the pharate adult (Shu et al., 1997). In non-Sphingid bombycoids, such as Bombyx mori (Bombycidae), Hyalophora cecropia (Saturniidae), and Malacosoma pluviale (Lasiocampidae), and in the lymantrid (Noctuoidea) Lymantria dispar, vitellogenesis proceeds before adult emergence in the absence of JH. In these groups, vitellogenesis seems totally independent of JH, as shown by allatectomy and JH treatment, which did not influence oocyte growth (see Wyatt and Davey, 1996 for references). Other studies of L. dispar, in which vitellogenesis starts as early as day 3 of the last larval instar, found that treatment with JHAs on day 2 of this instar selectively prevents the production of Vg (Fescemyer et al., 1992) and Vg mRNA (Hiremath and Jones, 1992). 3.9.2.1.4. Mosquitos and flies: the shift to ecdysteroid mediated vitellogenesis For the Diptera, edysteroids have the primary role of regulating reproduction, as demonstrated in detail almost exclusively in mosquitos and flies. JH plays an important priming role in these females by preparing reproductive tissues for ecdysteroid mediated processes. Regulation of Vg production by ecdysteroids was first demonstrated in the yellow fever mosquito, Aedes aegypti. Vg synthesis in the mosquito fat body is stimulated by a blood meal and inhibited by removal of ovaries prior to blood feeding (Hagedorn and Fallon, 1973). The ovaries secrete the factor required for Vg production by the fat body, and this was found to be ecdysone (E), which is converted into the active form of the hormone 20-hydroxyecdysone (20E) (Hagedorn et al., 1975). Remarkably, mosquitos and flies are the only insects known to use ecdysteroids as the key regulator of reproduction. How or why this shift away from JH- to ecdysteroid-mediated reproduction occurred in Diptera remains an enigma.

The endocrinology of vitellogenesis in mosquitos (suborder Nematocera) has been reviewed in detail for A. aegypti (Hagedorn, 1985; Raikhel, 1992a; Dhadialla and Raikhel, 1994; Sappington and Raikhel, 1998a; Raikhel et al., 2003; Wang et al., 2004) and investigated in Culex pipiens, Aedes atropalpus, and Anopheles stephensi (Hagedorn, 1985; Klowden, 1997). In A. aegypti, vitellogenesis is separated into four phases: previtellogenic (PV) preparation, arrest, yolk protein (YP) synthesis (vitellogenesis), and termination of vitellogenesis (Raikhel, 1992b; Dhadialla and Raikhel, 1994). A newly emerged female needs about 3 days to become competent for the physiological demands of intense vitellogenesis. During this phase, the fat body and ovary acquire responsiveness to 20E and become competent for blood meal-activated vitellogenesis and oogenesis, respectively (Flanagan et al., 1977; Li et al., 2000; Zhu et al., 2003a). After 3 days of PV preparation, the fat body and ovary enter a state of arrest that persists until a blood meal is taken. During the YP synthesis stage, proteins are produced by the fat body and accumulated by developing oocytes (Raikhel, 1992a; Raikhel et al., 2002). This massive synthesis peaks at around 24 h post-blood meal (PBM), then drops sharply, and terminates by 36–42 h PBM. The fat body undergoes remodeling, and the first batch of eggs completes chorion formation and is oviposited. Titers of JH III and ecdysteroids in female A. aegypti are presented in Figure 4. During PV preparation, JH III titer increases and stabilizes during the arrest phase, and the CA remains active (Shapiro et al., 1986). JH is required for the fat body to attain competence for YP synthesis in response to 20E. Following blood feeding, the JH III titer drops as a result of a rapid decrease in CA activity and elevation of JH esterase titer in the hemolymph (Shapiro et al., 1986). Blood feeding triggers the release of the ovarian ecdysteroidogenic hormone (OEH) from the medial neurosecretory cells of the brain for up to 12 h postfeeding (Figure 5; Lea, 1972; Brown et al., 1998). In response to OEH, the ovary produces ecdysteroids, and the hemolymph titer of ecdysteroids in female mosquitos is correlated with the rate of YP synthesis in the fat body (Hagedorn et al., 1975). The ecdysteroid titers are only slightly elevated at 4 h PBM, rise sharply at 6–8 h PBM to a maximum level at 18–20 h PBM, and then decline to previtellogenic levels by 30 h PBM (Hagedorn et al., 1975). Numerous studies have clearly established that ecdysteroid control of YP synthesis is a central event in the blood meal-activated regulatory cascade

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Figure 4 Juvenile hormone (JH) and 20-hydroxyecdysone titers and the level of Vg during the first reproductive cycle in the mosquito Aedes aegypti. (Based on data from Hagedorn, H., 1985. The role of ecdysteroids in reproduction. Kerkut, G.A., Gilbert, L.I. (Eds.), Comprehensive Insect Physiology, Biochemistry and Pharmacology, Vol. 8. Pergamon, Oxford, pp. 205–261; and Dhadialla, T., Raikhel, A.S., 1994. Endocrinology of mosquito vitellogenesis. In: Davey, K.G., Peter, R.E., Tobe, S.S. (Eds.), Perspectives in Comparative Endocrinology. National Research Council of Canada, Ottawa, pp. 275–281.)

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Figure 5 Schematic representation of the nutritional and hormonal activation of vitellogenesis in the mosquito Aedes aegypti. A blood meal results in a direct signal to the fat body, which is required to initiate yolk protein precursor (YPP ) gene expression. The brain also receives a signal from the midgut, which activates medial neurosecretory cells to release a peptide hormone, ovarian ecdysteroidogenic hormone (OEH). OEH activates follicular cells of the primary follicles to produce ecdysone. Ecdysone is converted in the fat body to the active steroid hormone, 20-hydroxyecdysone (20E), which activates YPP gene expression via an ecdysone hierarchy. Yolk protein precursors, vitellogenin (Vg), vitellogenic carboxypeptidase (VCP), vitellogenic cathepsin B (VCB), and lipophorin (Lp) are secreted by the fat body into the hemolymph and selectively accumulated by developing oocytes.

leading to successful egg maturation (reviews: Hagedorn, 1985; Raikhel, 1992b; Dhadialla and Raikhel, 1994; Wang et al., 2004). Consistent with the proposed role of 20E in activating mosquito YP

synthesis, experiments using fat body in vitro have shown that physiological doses of 20E (106 M) activate yolk protein precursor (YPP) genes, which are described below (Deitsch et al., 1995; Cho et al., 1999). Molecular studies have demonstrated that although the ecdysteroid triggered regulatory hierarchies, such as those implicated in the initiation of metamorphosis, are reiteratively utilized in the control of mosquito vitellogenesis, the unique interplay of hierarchical factors is determined by the mosquito’s biology (Raikhel et al., 2002, 2003; Wang et al., 2004). Of particular importance are the cyclicity of vitellogenic response and its dependence upon blood feeding. The demands for a very high level of expression of YPP genes, especially Vg, put additional restraints on the hormonal regulation of these genes. The endocrine control of vitellogenesis in higher flies (suborder Cyclorrhapha) has been studied extensively in several species (reviews: Bownes, 1986, 1994, 2004; Raikhel et al., 2003) and is altered according to whether the species lays eggs in batches or continuously. In species laying eggs in batches, the maturation of a synchronous group of oocytes is controlled by changes in hormone levels, as described best for the housefly, Musca domestica (Adams and Filipi, 1983; Adams et al., 1985). In this species, the fat body starts to synthesize YPs after the female fly feeds on a protein meal, and YP uptake also commences in the ovary. This phase is controlled by the circulating ecdysteroids, and the fat body later shuts down YP synthesis as egg development is completed. There is a strong correlation

440 Hormonal Control of Reproductive Processes

between ecdysteroid titers in the hemolymph and the progress of vitellogenesis, as well as between ecdysteroid titers and the amount of YP produced in the fat body (Adams et al., 1985). Ovaries produce ecdysteroids in vitro, and ovariectomized flies have reduced ecdysteroid levels early in the cycle (Adams et al., 1985). In the autogenic strain of M. domestica, decapitation of females blocks Vg gene expression in fat body, but 20E restores accumulation of Vg mRNA. Moreover, males were induced to produce YPs in response to the hormone (Agui et al., 1991). JH acts at an early stage to prime the fat body for YP synthesis and ensures that the oocytes are arrested in a previtellogenic stage (Adams, 1974, 1981). Interestingly, Adams et al. (1989) also have shown that the response to 20E in male houseflies was enhanced by application of JH. Many aspects of the regulation of vitellogenesis in the housefly are quite similar to those described for A. aegypti. In the blowfly, Calliphora vicina, YP-containing secretory granules become visible in the fat body when vitellogenesis is initiated by ingestion of a protein meal, but the granules are not detected in the fat body of ovariectomized females (Thomsen and Thomsen, 1974, 1978). Their appearance can be restored in ovariectomized females by injection of 20E (Thomsen et al., 1980). These observations have suggested the involvement of the ovary and ecdysteroids in the initiation of YP synthesis in the fat body of C. vicina and is reminiscent of the response of A. aegypti to a blood meal, which triggers the ovary to produce ecdysteroids and initiate a cycle of Vg production in the fat body. In the blowflies, Sarcophaga and Phormia, nonprotein-fed females respond to 20E by inducing YP synthesis in fat body (Huybrechts and De Loof, 1982). Unlike C. vicina, ovariectomized houseflies and blowflies still produce YPs indicating additional complexity of hormonal regulation in these insects (Engelmann and Wilkens, 1969; Jensen et al., 1981; Huybrechts and De Loof, 1982). Interestingly, male Sarcophaga, Phormia, and Lucilia also produce YPs in fat body in response to 20E injection (Huybrechts and De Loof, 1982). JH alone cannot induce YP synthesis in males in Musca or Calliphora (Adams, 1974; Huybrechts and De Loof, 1982; Adams et al., 1989). In those species continuously laying eggs, each individual egg chamber differentiates, and its progress through vitellogenesis is modulated by hormonal signals connecting its development to environmental factors such as mating and food intake. This type of vitellogenesis is characteristic

of D. melanogaster, and many related species. The regulation of vitellogenesis in D. melanogaster has been the subject of numerous studies using molecular, genetic, developmental, and physiological approaches (Bownes, 2004). In this species, adult sex determination (see Chapter 1.5) is the primary factor for correct expression of yolk protein genes. Once the genes are active in the female, the level of expression is modulated by ecdysteroids and JH in a complex way (Bownes et al., 1993; Bownes, 2004). JH seems crucial for vitellogenesis, especially in the ovary (Bownes et al., 1996; Soller et al., 1999), but it appears that JH does not directly affect the transcription of yp genes in the fat body. More recent data reported by Richard et al. (1998, 2001) points to a more prominent role for ecdysteroids in regulating vitellogenesis. It is interesting that in the Caribbean fruit fly, Anastrepha suspensa, in which the YPs are only produced in the ovary, there is no evidence of hormonal regulation by JH or 20E in females, and 20E injection is unable to stimulate YP synthesis in males (Handler, 1997). 3.9.2.2. Yolk Protein Precursors

3.9.2.2.1. Sites of yolk protein production In most insects, the fat body is the exclusive site for production of yolk protein precursors (YPPs) and, in others, the ovary is a complementary vitellogenic organ. Table 1 provides a list of species in which fat body and ovarian YPP production have been reported, and in cyclorrhaphan Diptera, YPP synthesis occurs in both the fat body and the ovary. In some dipteran species, such as the stable fly Stomoxys calcitrans, Anastrepha suspensa and perhaps the tsetse fly Glossina austeni, the ovary is the exclusive site of YPP synthesis (review: Wyatt and Davey, 1996). Insect YPs are the subject of several recent reviews (Bownes and Pathirana, 2002; Sappington et al., 2002; Telfer, 2002; Tufail et al., 2004; Masuda et al., 2004; Yamahama et al., 2004). 3.9.2.2.2. The fat body In female insects, the main function of the fat body is to produce massive amounts of YPPs. Multiple lobes of this organ are distributed mainly in the abdomen and to a lesser extent in the thorax and head. In many insects, fat body consists of a single cell type, the trophocyte (¼ adipocyte), and in cockroaches and some other insects, mycetocytes and urate cells are also present (Kelly, 1985; Locke, 1998). The multilobed structure of this tissue enhances its interaction with hemolymph. Also, the fat body is responsible for intermediary metabolism; storage of carbohydrates,

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t0005

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Table 1 Vitellogenic organs in insects Vitellogenic organ Order–suborder

Species

Fat body

Ovary

Type of ovarioles

Zygentoma Dictyoptera Dictyoptera Orthoptera–Caelifera Orthoptera–Ensifera Phasmida Hemiptera Coleoptera Coleoptera Hymenoptera Lepidoptera Lepidoptera Lepidoptera Diptera–Nematocera Diptera–Nematocera Diptera–Brachycera Diptera–Brachycera Diptera–Brachycera Diptera–Brachycera Diptera–Brachycera Diptera–Brachycera Diptera–Brachycera Diptera–Brachycera

Thermobia domesticaa Blattella germanica b Leucophaea maderae b Locusta migratoria b Acheta domesticusb Bacillus rossiusb Rhodnius prolixusc Leptinotarsa decemlineatab Coccinella septempunctata b Apis melliferad Plodia interpunctella b Hyalophora cecropia b Manduca sexta b Rhyncosciara americanab Aedes aegypti b Dacus oleaeb Drosophila melanogaster b Musca domestica b Sarcophaga bullata b Calliphora vicinab Stomoxys calcitrans b Anastrepha suspensa e Glossina austeni b

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

X

Panoistic Panoistic Panoistic Panoistic Panoistic Panoistic Telotrophic meroistic Telotrophic meroistic Telotrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic Polytrophic meroistic

?

X X X X

X X X X X X X

a

Rousset and Bitsch (1993). See Valle (1993) for references. c Melo et al. (2000). d K.R. Guidugli, M.D. Piulachs, X. Belles, and Z.L.P. Simo˜es, unpublished data. e Handler (1997). b

lipids, and proteins; and synthesis of hemolymph proteins. These functions are hormonally controlled and successively change in accordance with the demands of different life stages. Trophocytes are equipped with abundant cytoplasmic organelles that accomplish a great variety of synthetic and secretory functions. The basal lamina enveloping the fat body allows diffusion of large oligomeric proteins with molecular sizes of over 300 000 (even up to 500 000 Da) and keeps the apical plasma membrane of trophocytes structurally differentiated. This apical membrane is highly infolded and resembles a plasma membrane reticular system in which secretory granules are being released by exocytosis (Locke and Huie, 1983; Raikhel and Lea, 1983; Dean et al., 1985; Raikhel and Snigirevskaya, 1998; Mazzini et al., 1989; Giorgi et al., 2004). 3.9.2.2.3. Fat body derived yolk protein precursors 3.9.2.2.3.1. Vitellogenins In most female insects, the major constituent of protein yolk is Vg, a large, conjugated protein that is taken into oocytes and stored as vitellin (Vn). The amino acid sequence,

structure, and composition of Vg are sufficiently conserved between insects and other oviparous animals to indicate origin from a common ancestral protein (Chen et al., 1994; Sappington and Raikhel, 1998a; Sappington et al., 2002) and may share homology with other, more distantly related lipoproteins (Blumenthal and Zucker-Aprison, 1987; Spieth et al., 1991; Sappington et al., 2002). Insect Vgs are encoded by mRNAs of 6–7 kb that are translated as primary products of 200 kDa. Vg primary products have been characterized at the molecular level for Hemimetabola and Holometabola species (Sappington et al., 2002; Tufail et al., 2004). The primary pre-proVg is cleaved into subunits (apoproteins), ranging from 50 to 180 kDa, by subtilisin-like endoproteases and proprotein convertases (Barr, 1991; Rouille et al., 1995). These enzymes recognize a consensus motif (R/K)XX (R/K) preceding the cleavage site, and the motif is found in all known insect Vgs (Chen et al., 1994; Sappington and Raikhel, 1998a; Sappington et al., 2002; Tufail et al., 2004). Vg convertase (VC) has been characterized from the vitellogenic fat body of

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A. aegypti (Chen and Raikhel, 1996) and is a homolog of human and D. melanogaster furins and a D. melanogaster convertase (Barr et al., 1991; Roebroek et al., 1991, 1992a, 1992b; Hayflick et al., 1992). In hemimetabolous insects like L. maderae, P. americana, and Riptortus clavatus, the Vg primary product is cleaved into large and small polypeptides, including polypeptides of 80–110 kDa (Figure 6) (Della-Cioppa and Engelmann, 1987; Hirai et al., 1998; Tufail et al., 2001; Tufail and Takeda, 2002). It has also been reported that Vgs from some hemimetabolous insects, such as L. maderae and R. clavatus, are processed further in the oocyte (Hirai et al., 1998; Tufail and Takeda, 2002). In holometabolous insects, the Vg primary product is cleaved into a single large and small polypeptide (Figure 6) (Dhadialla and Raikhel, 1990; Chen et al., 1994, 1996). As with other apocritan

Hymenoptera, Vg is not cleaved in the parasitic wasp, Pimpla nipponica (Figure 6; Nose et al., 1997). In the fall armyworm moth, Spodoptera frugiperda, only a single Vg apoprotein was detected in hemolymph and ovarian extracts, although other lepidopteran Vgs normally are processed into two subunits (Hiremath and Lehtoma, 1997; Sorge et al., 2000). Following extensive co- and posttranslational modifications, Vg subunits form high molecular weight oligomeric phospholipoglycoproteins (400– 600 kDa) that are secreted into the hemolymph of females (Osir et al., 1986a; Wojchowski et al., 1986; Dhadialla and Raikhel, 1990; Sappington and Raikhel, 1998a; Giorgi et al., 1998; Tufail et al., 2004). Mature vitellogenins generally exist as oligomers, but monomeric molecules of about 300 kDa may exist in the cockroach N. cinerea (Imboden et al., 1987).

Figure 6 Schematic representation of the cleavage sites and polyserine domains in vitellogenins from 12 insect species. The arrows and white lines indicate the putative or determined cleavage sites following the consensus RXXR cleavage site sequence. The green segments show the polyserine stretches. Numbers indicate the amino acid residues deduced from the N-termini (excluding the signal peptides). Color code is used to indicate the number of vitellogenin subunits resulting from proteolytic cleavage. (Modified from Tufail, M., Raikhel, A.S., Takeda, M., 2004. Biosynthesis and processing of insect vitellogenins. In: Raikhel, A.S., Sappington, T.W. (Eds.), Reproductive Biology of Invertebrates, Vol. 12, Part B: Progress in Vitellogenesis. Science Publishers, Enfield, USA/Plymouth, UK.)

Hormonal Control of Reproductive Processes

One to several Vg genes have been identified in different insect species (reviews: Telfer, 2002; Tufail et al., 2004). In the silkworm B. mori, Vg is encoded by a single gene (Tufail et al., 2004), and there are five Vg genes in the mosquito A. aegypti (Romans et al., 1995). Regulatory regions of Vg genes that govern hormone dependent expression have been characterized in great detail for A. aegypti (Kokoza et al., 2001) and only partially for L. migratoria and B. germanica (Wyatt et al., 1984; Locke et al., 1987; Belles, 2004).

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3.9.2.2.3.2. Yolk polypeptides of the cyclorrhaphous Diptera In higher Diptera, the number of YPs ranges from one to five, and they are different from the Vg of other insects (review: Bownes and Pathirana, 2002). There are three major YPs of 46, 45, and 44 kDa (Barnett et al., 1980; Bownes et al., 1993) in D. melanogasti and up to five YPs ranging from 40 to 51 kDa in C. erythrocephala (Fourney et al., 1982), Neobellieria (¼ Sarcophaga) bullata (Huybrechts and De Loof, 1982), M. domestica (Adams and Filipi, 1983), A. suspensa (Handler, 1997), Phormia regina (Zou et al., 1988), Ceratitis capitata (Rina and Savakis, 1991), and eight other species of Drosophila (Bownes, 1980). The primary translation products of cyclorrhaphan YPs are close to the size of each mature polypeptide. Their posttranslational modification includes glycosylation and phosphorylation and tyrosine sulfation (Brennan et al., 1980; Minoo and Postlethwait, 1985; Baeuerle and Huttner, 1985). Yolk protein genes have been characterized for D. melanogaster (Hung and Wensink, 1983; Garabedian et al., 1987; Yan et al., 1987), C. capitata (Rina and Savakis, 1991), and C. erythrocephala (Martinez and Bownes, 1994). Deduced amino acid sequences for the three YPs of D. melanogaster show that they are related to each other and to other cyclorrhaphan YPs. These YPs are not lipoproteins (see Chapter 4.6), likely constitute subunits of a larger native protein (Fourney et al., 1982; Adams and Filipi, 1983; Zou et al., 1988), and more closely resemble a family of vertebrate lipases and not the insect Vgs (Bownes et al., 1988; Terpstra and Ab, 1988). 3.9.2.2.3.3. Additional yolk proteins secreted by the fat body A number of supplemental proteins are secreted by the fat body of vitellogenic females and selectively accumulated by developing oocytes. Typically, these YPs are minor yolk components but in some species can be as abundant as Vg (Telfer, 2002; Masuda et al., 2004). Most are female-specific products, but some are found in the hemolymph

443

of both sexes. They are likely to serve specialized functions necessary for embryonic development and, as yet, it is difficult to draw any unifying conclusions, since very few of these YPs have been characterized for insects. 3.9.2.2.3.3.1. Microvitellogenin Microvitellogenin (mVg) is a small yolk protein of 30 kDa (Pan, 1971; Kawooya et al., 1986; Telfer and Pan, 1988; Pan et al., 1994). In lepidopteran species, mVg is synthesized and secreted by the fat body (Cole et al., 1987) and incorporated by ovarian follicles (Telfer and Kulakosky, 1984; Kulakosky and Telfer, 1989). Manduca sexta mVg is a monomeric protein with no detectable carbohydrate, lipid, or phosphate (Kawooya et al., 1986). Bombyx mori produces multiple 30 kDa proteins that are the principal components of both male and female hemolymph during late larval and pupal stages (Izumi et al., 1981) and constitute 35% of the egg total soluble protein (Zhu et al., 1986). Deduced amino acid sequences from five B. mori mVg cDNA clones revealed their high similarity (Sakai et al., 1988), and M. sexta mVg shares 70% sequence similarity with a B. mori mVg (Wang et al., 1989). Furthermore, antigenic similarity between mVgs in M. sexta and H. cecropia (Kawooya et al., 1986) supports the homology of lepidopteran mVgs. 3.9.2.2.3.3.2. Lipophorin as a yolk protein Lipids are a critical source of energy during insect embryogenesis (Beenakkers et al., 1985) and can represent as much as 30–40% of the egg’s dry weight (Troy et al., 1975; Kawooya and Law, 1988; Briegel, 1990). Lipophorins (Lp) are lipid transport proteins in insects (see Chapter 4.6) and play a prominent role in the accumulation of lipids in insect oocytes (review: Antwerpen van et al., 2004). Kinetic analyses have indicated that lipid transfer is affected by a saturable mechanism in both M. sexta (Kawooya and Law, 1988) and H. cecropia (Kulakosky and Telfer, 1990), thus indicating that lipid uptake occurs via receptor-mediated endocytosis. Specific ovarian receptors for Lp have been cloned from L. migratoria (Dantuma et al., 1997), A. aegypti (Cheon et al., 2001), and B. germanica (Ciudad, Piulachs, and Belles, unpublished data). In saturniid and sphingid moths, Lp is the second most abundant YP (Chino et al., 1977b; Telfer et al., 1991; Telfer and Pan, 1988), and it has been found in the yolk of the fall webworm H. cunea (Arctiidae) (Yun et al., 1994). In mosquito eggs, Lp makes up only 3% of total egg proteins (Sun et al., 2001). In B. germanica, Lp facilitates hydrocarbon uptake by maturing oocytes (Fan et al., 2002).

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3.9.2.2.3.3.3. Pro-proteases Yolk proteins are essential reserves of amino acid and other nutrients for the developing insect embryos, and sequential cleavage of YPs into smaller molecules occurs throughout embryogenesis (Zhu et al., 1986; Yamashita and Indrasith, 1988; Masetti and Giorgi, 1989; Yamamoto and Takahashi, 1993; Izumi et al., 1994; Takahashi et al., 1996; Cho et al., 1999; Yamahama et al., 2004). This proteolytic degradation of YPs is probably regulated through a battery of proteases (see Chapter 4.7). At present, two female-specific proenzymes are known to be deposited in the protein yolk of A. aegypti. One is a serine carboxypeptidase, vitellogenic carboxypeptidase (VCP), and it is a glycosylated 53 kDa protein secreted by the female fat body in synchrony with vitellogenin (Hays and Raikhel, 1990). An antigenically similar protein of the same size occurs in the yolk, and the enzyme is activated during embryogenesis by cleavage to a 48 kDa polypeptide (Cho et al., 1991). A 44 kDa fat body protein, vitellogenic cathepsin B (VCB), with sequence similarity to vertebrate cathepsin B is also incorporated into the yolk of A. aegypti (Cho et al., 1999). It is converted to 42 kDa after internalization in developing oocytes, and then to 33 kDa in developing embryos, coinciding with the onset of YP degradation. The 33 kDa protein degrades mosquito YPs in vitro. Secretory pathways for VCP and VCB in the fat body and their endocytosis in the oocyte were shown by immunocytochemistry to be the same as those of vitellogenin (Snigirevskaya et al., 1997a). Notably, they are deposited in the amorphous, peripheral layer of the yolk bodies surrounding the central core of the vitellin crystal (Snigirevskaya et al., 1997b). An acid cysteine proteinase (see Chapter 4.7) was first purified from Bombyx eggs (Kageyama and Takahashi, 1990) and named Bombyx cysteine proteinase (BCP). It has broad substrate specificity and hydrolyzes various protein substrates, including B. mori yolk proteins. Since BCP accumulates in hemolymph, it is secreted by the fat body as shown by Northern blots. Ovarian follicle cells also synthesize the enzyme, as established by Northern blot and its synthesis from ovarian RNA (Yamamoto et al., 2000). 3.9.2.2.3.3.4. Other fat body products used as yolk proteins For several insects, there are reports of other proteins that are incorporated into yolk. In L. migratoria females, 21 kDa and 25 kDa proteins are produced by the fat body in synchrony with the synthesis of Vg (Zhang et al., 1993; Zhang and

Wyatt, 1996) and accumulate in oocytes along with Vg. These additional yolk proteins are significant constituents of the yolk in L. migratoria eggs. In the lepidopteran M. sexta, a blue biliprotein (insecticyanin) composed of four 21.4 kDa subunits is a component of larval hemolymph and is present in eggs (Chino et al., 1969; Kang et al., 1995). An arylphorin-like cyanoprotein that is a hemolymph storage hexamer is also deposited in eggs of the bean bug R. clavatus (Chinzei et al., 1990; Miura et al., 1994). The incorporation of these pigmented proteins may help to conceal eggs from predators (Chino et al., 1969). The iron transport protein, transferrin (see Chapter 4.10) has been shown to be selectively deposited in yolk of the flesh fly Sarcophaga peregrina (Kurama et al., 1995) and the bean bug R. clavatus (Hirai et al., 2000). Hemolymph and oocytes of Rhodnius prolixus also contain a 15 kDa heme binding protein (Oliveira et al., 1995). Eggs of the stick insect C. morosus contain a minor yolk protein that is secreted by the fat body and sulfated by the follicle cells (Giorgi et al., 1995). 3.9.2.2.4. Yolk proteins synthesized by ovarian follicle cells Irrespective of insect ovary type, only the follicle cells in egg chambers engage in the production of proteins that are utilized as YPs. As demonstrated for several insects, follicle cells secrete a protein that has a similar antigenic reactivity and subunit composition as the Vg produced by fat body. In the firebrat T. domestica, there is the dual origin of YP from fat body and ovaries (Rousset and Bitsch, 1993) (Table 1). The synthesis of YPs by both tissues in adult females of this order, where molting and reproduction alternate, indicates that it could be an ancestral condition. YP production has also been observed in ovarian follicles of the heteropteran R. prolixus (Melo et al., 2000) and two coleopterans, C. septempunctata (Zhai et al., 1984) and L. decemlineata (Peferoen and De Loof, 1986). In the honeybee, A. mellifera, Vg expression has been reported in the fat body (Piulachs et al., 2003), but a more recent study has revealed that it is also expressed in the ovaries (K.R. Guidugli, M.D. Piulachs, X. Belles, and Z.L.P. Simo˜ es, unpublished data). These findings in R. prolixus and A. mellifera suggest the importance of reassessing the vitellogenic role of the ovary in more species. In higher flies (suborder Cyclorrhapha), the ovarian origin of YPs was first detected in D. melanogaster (Bownes and Hames, 1978), and in this and other species, these YPs have been investigated extensively (see Section 3.9.2.2.3.2). Follicle cells were identified as the site of synthesis in D. melanogaster

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by their ability to secrete YPs when manually separated from follicles labeled with [35S]methionine, and by in situ hybridization to contain YP mRNAs (Brennan et al., 1982). Follicle cells are implicated in production of YPs in C. erythrocephala (Rubacha et al., 1988) and in S. bullata (Geysen et al., 1986). In the stable fly, S. calcitrans (Houseman and Morrison, 1986; Chen et al., 1987), the Caribbean fruit fly A. suspensa (Handler and Shirk, 1988; Handler, 1997), and the tsetse fly, glossina austeni (Huebner et al., 1975), female specific proteins do not occur in the hemolymph of reproductive adults, thus indicating that YPs are synthesized only in the ovaries. In some cyclorrphan flies, YP genes are expressed differently in the fat body and follicular epithelium. In C. erythrocephala, ovaries produced 51 and 49 kDa YPs, while fat body secreted a 46 kDa YP and possibly a different 49 kDa YP (Fourney et al., 1982). In D. melanogaster, the two tissues secrete the same three YPs, but the smallest one,YP3, is underrepresented in ovarian synthesis (Brennan et al., 1982; Isaac and Bownes, 1982). Ovarian and fat body YPs are mixed in the eggs of these flies. In several species of Lepidoptera, follicle cells secrete proteins that are incorporated into yolk and may account for up to 25% of the total soluble protein, as in mature eggs of B. mori (Zhu et al., 1986). In the moth H. cecropia, a 55 kDa protein is present in the intercellular spaces and yolk bodies (Bast and Telfer, 1976). Follicles of B. mori produce an ovary specific protein of 225 kDa with three 72 kDa subunits (Ono et al., 1975; Indrasith et al., 1988; Sato and Yamashita, 1991a); one of which is converted to a 64 kDa polypeptide by egg maturation. Sequencing of a cDNA clone revealed its similarities to human gastric and rat lingual lipases, especially to a noncatalytic lipid binding domain (Inagaki and Yamashita, 1989; Sato and Yamashita, 1991b). Two follicle specific YPs have also been isolated from M. sexta (Tsuchida et al., 1992). One is a 130 kDa protein consisting of two glycosylated and phosphorylated 65 kDa subunits, similar to those of the follicle cell derived yolk protein of B. mori. The second is a slightly larger 140 kDa protein that is glycosylated but not phosphorylated. In the pyralid moth P. interpunctella, two yolk polypeptides originate in the follicle cells, and the 235– 264 kDa protein with subunits of 69 and 33 kDa is an ortholog of the egg specific protein in B. mori (Shirk et al., 1984; Bean et al., 1988; Shirk and Perera, 1998). Sequences for a cDNA clone encoding YP4 of P. interpunctella (Perera and Shirk, 1999) showed it to be an ortholog of a follicular product in G. mellonella (Rajaratnam, 1996).

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3.9.2.3. Mechanisms of Juvenile Hormone Action in Vitellogenesis

3.9.2.3.1. Induction of vitellogenin gene expression Vitellogenic action of JH is one of the hallmarks of insect reproduction. Although JH is known to initiate vitellogenesis in many insects (see Chapter 3.7), its precise mechanism of action is poorly understood. Studies of the induction of Vg gene transcription by JH have focused on two species: the migratory locust, L. migratoria, and the German cockroach, B. germanica (Wyatt, 1991, 1997; Belles, 2004). Two Vg genes have been partially characterized in L. migratoria, and studies at the molecular level have demonstrated the absence of detectable Vg mRNA in the fat body of JH deprived specimens (Dhadialla et al., 1987). Transcription of both Vg genes is coordinately induced in vivo by JH or JHA (Dhadialla et al., 1987; Glinka and Wyatt, 1996), but allatectomized locust females have a low sensitivity to JH, since doses up to 100 mg of JH III do not induce Vg production. JHAs, such as methoprene or pyriproxyfen, are more potent and stable (Wyatt et al., 1996). When JHA is administered to females chemically allatectomized with the allatocide precocene within 1 day of adult emergence, there is a lag of 12–24 h before Vg mRNA can be detected (Edwards et al., 1993). This lag period can be shortened by prior administration of a subeffective dose of JH or JHA that is insufficient by itself to induce Vg gene transcription. This dose likely primes the fat body cells for an accelerated response to a subsequent effective dose (Figure 7) (Edwards et al., 1993; Wyatt et al., 1996). The lag period between JHA treatment and vitellogenesis can be extended by inhibition of protein synthesis with cycloheximide (Figure 7) (Edwards et al., 1993). These results suggest that the action of JH on Vg genes in locust females is indirect and requires the synthesis of protein factors involved in transcription. In this respect, the priming action of JH on fat body for a vitellogenic response is similar to that in mosquitos in which the molecular nature of this JH action has been recently elucidated (Zhu et al., 2003a). Studies on the regulation of vitellogenesis in B. germanica also were facilitated by the cloning of a Vg cDNA (Martin et al., 1998; Comas et al., 2000). Experiments in vivo with allatectomized females have shown that Vg mRNA can be detected as early as 2 h after treatment with 1 mg of JH III, whereas Vg protein can be detected 2 h later (Figure 8). In addition, dose–response studies show that doses of 0.1, 1, and 10 mg of JH III induced Vg

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Figure 7 Induction of vitellogenin transcription in chemically allatectomized females of the migratory locust, Locusta migratoria. (a) Shortening of the response lag time by administration of a subeffective dose of JH. Specimens pretreated with low doses of JH III in acetone (four applications of 10 mg each, over 48 h) and then treated with 10 mg of pyriproxyfen, synthesize vitellogenin earlier (circles) than those receiving an equivalent treatment of acetone alone and pyriproxyfen (triangles). Equivalent pretreatment with JH III, but no pyriproxyfen, did not induce detectable vitellogenin synthesis (squares). (Data from Wyatt, G.R., Braun, R.P., Zhang, J., 1996. Priming effect in gene activation by juvenile hormone in locust fat body. Arch. Insect Biochem. Physiol. 32, 633–640.) (b) Inhibitory effects of cycloheximide (CHX) upon vitellogenesis induced by pyriproxyfen. Insects were treated with cycloheximide (62 mg) in water or with water alone, and 1 h later treated with 10 mg pyriproxyfen. Cycloheximide delayed vitellogenin transcription by about 1 day, which is approximately equal to the duration of inhibition of protein synthesis. (Reproduced with permission from Edwards, G.C., Braun, R.P., Wyatt, G.R., 1993. Induction of vitellogenin synthesis in Locusta migratoria by the juvenile hormone analog, Pyriproxyfen. J. Insect Physiol. 39, 609–614.)

synthesis, but not 0.01 mg (Comas et al., 1999, 2001). Cycloheximide applied to fat body in vitro abolishes the vitellogenic effects of JH (Comas et al., 2001), again suggesting that the effect of JH on the Vg gene involves the synthesis of protein factors involved in transcription.

Figure 8 Production of vitellogenin mRNA and vitellogenin protein in vivo after JH treatment of allatectomized females of Blattella germanica. A dose of 1 mg of JH III was topically applied to 48 h old allatectomized females; the fat body was dissected 2, 4, 6, 8, or 10 h later, and analyzed for vitellogenin mRNA (Northern blot, above), or vitellogenin protein (Western blot, below). (Reprinted with permission from Comas, D., Piulachs, M.D., Belles, X., 1999. Fast induction of vitellogenin gene expression by juvenile hormone III in the cockroach Blattella germanica (L.) (Dictyoptera, Blattellidae). Insect Biochem. Mol. Biol. 29, 821–827, with permission from Elsevier.)

3.9.2.3.2. Potential response elements in JHdependent genes related to vitellogenesis Results from the above studies of JH action on vitellogenesis in the locust and cockroach led to the hypothesis that JH may affect proteins belonging to the nuclear hormone receptor superfamily (see Chapters 3.5 and 3.6), which in turn would mediate transcription of Vg genes (Wyatt et al.,1996; Belles, 2004; see Chapter 3.5). Signature response elements for the binding of the putative JH transcription factors could be found within the regulatory regions of these genes, in the same way as those for the ecdysteroid receptor (EcR) and related proteins. Analysis of the jhp21 gene of L. migratoria revealed the partially palindromic, 13-nucleotide motif AGGTTCGAGA/TCCT that is found in three copies from the transcription start point (Zhang and Wyatt, 1996). This motif is suggestive of a hormone response element, given that it is similar to the consensus ecdysteroid response element (see Chapter 3.5), as defined by Jiang et al. (2000). Furthermore, this nucleotide motif is very similar to the canonical sequence IR-1 (AGGTCAATGACCT), a

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consensus-inverted repeat with a single nucleotide spacer that is recognized by the ecdysteroid receptor and that confers JH responsiveness upon genes in mammalian cells that contain farnesoid x receptor (FXR), a member of the nuclear receptor superfamily (Forman et al., 1995).

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3.9.2.3.3. Molecular mechanisms in juvenile hormone action Hypothesized JH specific DNA motifs require functional characterization, either by demonstrated activity in gene regulation or by binding to a known JH receptor or a transcription factor involved in JH response. Significant progress in this regard has been made on the 13-nucleotide motif AGGTTCGAGA/TCCT from the upstream region of the jhp21 gene in L. migratoria. A cellfree transcription system was developed that uses nuclear proteins extracted from locust fat body, and transcription is measured with reporter constructs containing a short DNA sequence, lacking G in the transcribed strand, fused to the promoter sequence of interest (Zhang et al., 1996). When transcription is carried out with the transcription terminator O-methyl-GTP and not GTP, only the DNA sequence lacking G is transcribed, and the transcript can be resolved by gel electrophoresis. With nuclear extracts from untreated adult female L. migratoria, constructs that include the promoter region of the vitellogenin or jhp21 genes are transcribed, as is also the nonspecific promoter of the adenovirus major late antigen (AdML), which is used as a positive control. However, extracts from precocene treated females, while still transcribing AdML, fail to transcribe from the jhp21 promoter (Zhang et al., 1996). After transcription was found to be specific for the reproductive female fat body, truncated constructs of the promoter region of the jhp21 gene were used to show that the DNA between nucleotides 1056 and 1200 from the transcription start site strongly enhanced transcription. In synthetic constructs, the incorporation of two tandem copies of the 15 nucleotide element GAGGTTCGAGACCTC (found at 1152) stimulated transcription as strongly as the native 145 nucleotide sequence, whereas the 15 nucleotide element, mutated at four positions and inserted into two copies, was inactive (Zhang et al., 1996). These results suggested that the sequence GAGGTTCGAGACCTC might be a JH response element. Tests of nuclear extracts for specific protein binding to the putative JH response element with the electrophoretic mobility shift assay demonstrated the occurrence of a specific DNA binding protein in extracts from JH exposed fat body, whereas it was found to be lacking in JH deprived tissue (Zhang et al., 1996). Furthermore, Zhou et al. (2002) have reported

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that this binding shows a preference for the inverted repeat GAGGTTC in the left half-site and that it is abolished by phosphorylation catalyzed by a protein kinase C present in the nuclear extracts. These results further support the identification of a putative JH response element that is bound by a transcription factor brought to an active state by JH. It is still uncertain whether the binding protein may be the nuclear JH receptor, a dimerization partner of the receptor, or another protein factor involved in the transcriptional process (Wyatt, 1997). A putative JH response element has been identified in the JH esterase gene (Cfjhe) from the spruce budworm, C. fumiferana (Kethidi et al., 2004). This 30 bp region contains two conserved hormone response element half-sites separated by a 4 nucleotide spacer similar to the direct repeat 4 (DR-4). The response element designated as JHRE is located between 604 and 574 of the Cfjhe promoter and is sufficient to support JH I induction. At the same time, the same region is responsive for 20E mediated repression of this gene. When a luciferase reporter was placed under the control of JHRE, a minimal promoter was induced by JH I in a dose and time dependent manner in a cell transfection assay. Moreover, the gel-retardation assay revealed the presence of a JHRE binding protein in the nuclear extract isolated from JH I treated CF-203 cells. The results described in the following sections suggest the hypothesis that JH acts via a nuclear receptor mode of action at the transcriptional level. 3.9.2.3.4. Is there a juvenile hormone nuclear receptor? Early experiments studying farnesol related molecules as possible ligands for the retinoid x receptor (RXR)–FXR mammalian receptor complex showed that JH III activated this complex (although it did not activate FXR or RXR alone), whereas methoprene did not (Forman et al., 1995). Curiously enough, an independent study reported that RXR alone could be activated by methoprene acid but not by JH III (Harmon et al., 1995). Although the physiological significance of these experiments remained unclear, attention has been drawn to Ultraspiracle (USP) (insect ortholog of RXR and obligatory dimerization partner for the EcR) as a possible candidate for a JH receptor. Work along this line was published by Jones and Sharp (1997), who reported that JH at micromolar concentrations binds to D. melanogaster USP, modifying its conformation and inducing USP dependent transcription, whereas yeast two-hybrid assays indicated that JH could promote USP homodimerization (see Chapters 3.5 and 3.7). In response to ligand binding, D. melanogaster USP undergoes

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conformational change to form a multihelix hydrophobic groove for recruitment of transcriptional coactivators (Jones and Jones, 2000). In addition, JH III binds to the ligand binding pocket of USP, and application of JH III to cells activates a transfected reporter construct containing a JH esterase core promoter and a DR12 hormone response element. The DR12 element confers enhanced transcriptional JH responsiveness, and it binds with specificity to recombinant USP (Jones et al., 2001; Xu et al., 2002). In more primitive insects, USP has a ligand binding domain closer to vertebrate RXR than to dipteran or lepidopteran USP (Bonneton et al., 2003). This has raised the question of whether an RXR ortholog of primitive insects might bind JH with higher affinity than USPs of higher insects. This possibility has been explored in L. migratoria, in which two isoforms of RXR are known: LmRXRL is more closely related to vertebrate RXR and LmRXR-S has a deletion of 66 nucleotides in the ligand binding domain (Hayward et al., 2003). Both LmRXR-S and LmRXR-L formed heterodimers with locust LmEcR in vitro, which bound the active ecdysteroid ponasterone A, as expected. However, neither LmRXR isoform alone nor LmRXR heterodimerizing with LmEcR bound JH III at nanomolar concentrations (Hayward et al., 2003). Parallel work in B. germanica has led to cloning of the orthologs BgRXR-L and BgRXR-S. Interestingly, despite the highly fluctuating levels of circulating JH III, mRNA levels of both BgRXR-L and BgRXR-S isoforms in the fat body remain constant throughout the vitellogenic cycle (Maestro, Martin and Belles, unpublished data). If either insect USP or RXR is a JH receptor, it behaves in a very different way to other members of the steroid/thyroid hormone nuclear receptor family, especially since its binding of JH is not saturable and is of relatively low affinity. One possibility is that USP or RXR form a heterodimer with a nuclear receptor other than EcR and that this complex might play the role of a JH receptor. Another possibility is inspired by promiscuous nuclear receptors like RXR, PPAR, or PXR of vertebrates (Harmon et al., 1995; Zomer et al., 2000; Watkins et al., 2001), which are activated by a variety of effectors and have Kd values in the micromolar range. These nuclear receptors have been considered as chemical sensors rather than canonical specific receptors (Watkins et al., 2001), a concept that could be extended to the interaction of JHs with the RXR/USP of insects. The Methoprene resistant (Met) gene of Drosophila may also be a useful model for gaining insight

into the molecular action of JH. Mutant D. melanogaster resistant to methoprene were generated in the 1980s, and these Met mutants were also highly resistant to JH III, JH bisexpoxide (JHB3), and other JHAs, but not to conventional insecticides. Biochemical work later showed the presence of an 85 kDa protein in various tissues of wild-type D. melanogaster that bound specifically to JH III with high affinity (Kd ¼ 6.7 nM), whereas in Met mutants the affinity of this protein for the hormone was lower by six-fold. In addition, JH III stimulation of protein synthesis in male accessory glands was clearly less pronounced in Met mutants than in wild-type flies, which suggested that Met is involved in the mechanism of action of JH (Shemshedini and Wilson, 1990; Shemshedini et al., 1990; see Chapter 3.7). Sequencing of the corresponding gene showed that Met is a member of the bHLH-PAS family of proteins that behave as transcriptional regulators (Ashok et al., 1998). These proteins show a basic helix–loop–helix domain (bHLH), which is characteristic of a number of transcription factors, and a PAS domain, named after the first members of the family: the products of the genes period (Per) (see Chapter 4.11) and single-minded (Sim) from D. melanogaster, and the aryl hydrocarbon receptor (AhR) and the Ah receptor nuclear translocator (ARNT) from vertebrates. Interestingly, Per and Sim have dimerization partners but no known ligands. AhR and ARNT form a functional dimer in the presence of a ligand, which upregulates a series of genes, the products of which metabolize foreign chemicals. Although Met mutants show no serious problems during embryonic and larval development or during metamorphosis (Wilson and Ashok, 1998), it is tempting to consider that the Met product could be involved in the transcription of JH-target genes, either by being the JH receptor or a transcription factor of the cascade triggered by JH. These results also support the hypothesis that Met could be a receptor coactivator for the JH receptor, playing a role similar to that reported for the steroid hormone coactivator-1, the disruption of which results in hormone resistance in mice (Xu et al., 1998). In this context, it is worth noting that the steroid hormone coactivator-1 also belongs to the bHLH-PAS family and that it and Met have the LXXLL motif, which in vertebrate transcriptional activators is needed for binding to nuclear receptors. The identification of JH response elements in JH responsive genes might be the most promising approach to the characterization of the JH receptor. This strategy is being followed for the jhp21 gene in L. migratoria and for the cfihe gene in

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C. fumiferana, with the encouraging results described above. Also promising has been the discovery of JH responsive genes in D. melanogaster, such as JhI-1, JhI-21, JhI-26, and minidiscs (mnd) (Dubrovsky, 2002), whose promoter regions may provide the needed JH response elements more easily, given the genetic, conceptual, and technical background available in studies of this fly. Along this line, promoter regions of the JhI-26 and mnd genes, which are directly inducible by JH, have already been tested for their ability to confer JH inducibility on heterologous reporter genes transfected to S2 cells. Constructs prepared with a 3 kb fragment from a region upstream of either the JhI26 or the mnd transcription initiation site cloned in front of the hsp70-lacZ reporter gene have served for the first experiments. After transfection and methoprene induction, the JhI-26 promoter insertion conferred between a 6- and 15-fold increase in the 8-galactosidase activity, whereas that of mnd conferred between a three- and fourfold increase (Dubrovsky, 2002). The results suggest that these 3 kb promoter fragments contain JH response elements, and thus further study of shorter fragments should lead to the determination of their role. In the meantime, in silico studies on the promoter fragments used in the transfections have revealed several motifs showing a significant similarity with the estrogen response element (ERE), suggesting that they can be considered putative response elements (Dubrovsky, 2002). 3.9.2.4. Molecular Mechanisms of Ecdysteroid Action in Vitellogenesis p9000

The early 20E-inducible gene E75 has been implicated in the JH signaling pathway (Dubrovsky et al., 2004). JH induces the E75A orphan nuclear receptor in D. melanogaster. The induction by JH is rapid and does not require protein synthesis, indicating direct action of JH on the E75 gene. E75A mRNA is reduced in ovaries of apterous4 Drosophila mutant adults defective in JH secretion. However, E75A mRNA expression can be rescued by topical JH analog application. Furthermore, ectopic expression of E75A is sufficient to perform several functions of the JH signaling pathway. Ectopic E75 can downregulate its own transcription and potentiate the JH inducibility in the JH gene, JhI-21. In the presence of JH, E75A can also repress 20E activation of early genes including Broad. The occurrence of a putative response element for E75 (the motif TGACCAAATT) (Belles, 2004) in the promoter region of the Vg gene of B. germanica, led to clone the three isoforms of this transcription factor, BgE75A, BgE75B, and BgE75C, in this cockroach.

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The pattern of expression of the three isoforms and results of RNAi experiments suggest that JH enhances E75 induction by 20E and that E75 is involved in the modulation of the cycles of oocyte maturation in B. germanica (D. Man˜ e´ , D. Martin and X. Belles, unpublished). In contrast to JH, the molecular mechanisms governing 20E regulation of reproductive events have been elucidated in detail for the yellow fever mosquito, A. aegypti, and the fruit fly, D. melanogaster (see Chapters 3.5 and 3.6). Owing to variations in their reproductive biology, studies of these insects often cover different events in female reproductive endocrinology, thus providing complementary information. 3.9.2.4.1. Genetic regulation of ecdysteroid response Ashburner et al. (1974) proposed a hierarchical model for the genetic regulation of polytene chromosome puffing by 20E (see Chapter 3.5). According to this model, the binding of 20E to its cognate receptor triggers the expression of a small set of early genes, which in turn activate expression of a large set of late genes and which, meanwhile, repress their own transcription. Subsequent genetic and molecular biological studies have revealed events underlying this ecdysteroid regulatory hierarchy in D. melanogaster (Thummel, 1996, 2002; Bender, 2003). Responses to ecdysteroids are mediated by the EcR complex, which consists of two members of the nuclear receptor gene family, EcR and USP, the latter being the retinoid X receptor homolog (Koelle et al., 1991; Yao et al., 1992, 1993). Upon binding 20E, this heterodimer recognizes a sequence specific DNA motif, ecdysteroid response element (EcRE), and directly induces the transcription of a small set of so-called early genes. These early genes encode mainly transcription factors, including E74, E75, and Broad-Complex (BR-C), of the Ets, nuclear receptor, and zinc-finger type, respectively, that transfer and amplify the hormonal signal through the regulation of numerous late ecdysone-responsive genes that specify stage and tissue specific effects of 20E (Thummel, 1996, 2002). The E74, E75, and BR-C early genes each encode a family of transcription factors by use of multiple promoters and alternative splicing. Biochemical and mutational analysis has shown that each of these genes acts as a 20E induced transcription factor and is required for the hormonal response in target tissues (Thummel, 2002; Bender, 2003). 3.9.2.4.2. Ecdysteroid receptor (EcR) The diversity of cellular responses to 20E may require particular combinations of EcR isoforms. Indeed,

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Figure 9 Expression profiles and hormone responsiveness of the key factors of the 20-hydroxyecdysone (20E) regulatory hierarchy and their target genes, yolk protein precursors (YPP ), during vitellogenesis in Aedes aegypti. Hormone titers of 20E in A. aegypti females (Hagedorn et al., 1975; Shapiro et al., 1986). Transcript profiles and sensitivities of the transcription factors to 20E (AaEcRA and AaEcRB, AaUSPA and AaUSPB, AaE74A and AaE74B, and AaHR3) and the late genes (Vg, VCP, and VCB ) were determined by Northern or RT-PCR analyses. Adapted from Cho and Raikhel (1992; Vg), Cho et al. (1991; VCP ), Cho et al. (1999; VCB), Wang et al. (2002; AaEcRA and AaEcRB), Wang et al. (2000b; AaUSPB and the 20E repressible AaUSPA), Sun et al. (2002; AaE74A and AaE74B), Pierceall et al. (1999; AaE75A), Kapitskaya et al. (2000; AaHR3), and L. Chen, J. Zhu, and A.S. Raikhel (unpublished data; BR-C isoforms Z2 and Z4). E, eclosion; BM, blood meal. (Modified with permission from Sun, G.Q., Zhu, J.S., Raikhel, A.S., 2004. The early gene E74B isoform is a transcriptional activator of the ecdysteriod regulatory hierarchy in mosquito vitellogenesis. Mol. Cell. Endocrinol. 218, 95–105.)

expression of specific D. melanogaster EcR isoforms could be correlated with patterns of responses of particular cell or tissue types to 20E (Talbot et al., 1993). Genetic studies have confirmed that different EcR proteins are functionally distinct, with the greatest difference being between the EcR-A and EcR-B isoforms and the greatest overlap being between the EcR-B1 and EcR-B2 functions (Bender et al., 1997; Schubiger et al., 1998; Lee et al., 2000; see Chapter 3.5). Cloning of the A. aegypti EcR isoforms, AaEcR-A and AaEcR-B, has facilitated evaluation of their expression during mosquito vitellogenesis. Significantly, transcripts of both isoforms exhibit different patterns of expression after a blood meal triggers fat body vitellogenesis in female mosquitos (Figure 9). The AaEcR-B transcript level rose sharply by 4 h PBM, which coincides with the small ecdysteroid

peak, and then declined and reached its lowest level at 16–24 h PBM. In contrast, the AaEcR-A transcript peaked at 16–20 h PBM, coinciding with the large ecdysteroid peak (Cho et al., 1995; Wang et al., 2002). Both isoform mRNAs were transcribed in a cycloheximide independent manner, suggesting that they are direct targets of 20E. However, AaEcR-A transcription requires the continuous presence of 20E, while the AaEcR-B mRNA level rose for 4 h and then declined under the same conditions. These results indicate that the mosquito EcR isoforms play distinct physiological functions during vitellogenesis in the mosquito fat body. 3.9.2.4.3. USP isoforms Like the EcR isoforms, two USP cDNA isoforms (AaUSP-A and AaUSP-B) were cloned from A. aegpyti (Kapitskaya et al., 1996). They differ only at the N-terminus, thus

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indicating their origin from the same gene, which recently has been confirmed by cloning the USP gene from A. aegypti (Wang, Zhu, and Raikhel, unpublished data). The isoform specific expression patterns in the Aedes fat body during vitellogenesis suggest that the two mosquito USP isoforms may carry out distinct functions. The level of AaUSP-B mRNA correlates with the high titer of ecdysteroid at 16–20 h PBM (Wang et al., 2000b). Intriguingly, USP-B is activated with 20E, whereas USP-A is inhibited by the hormone (Figure 9) (Wang et al., 2000b). AaEcR heterodimerizes with either mosquito USP isoform; the AaEcR-USP-B heterodimer displays stronger DNA binding and transactivation activities than the AaEcR-USP-A heterodimer (Wang et al., 2000b).

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3.9.2.4.4. Ecdysteroid response elements (EcREs) The canonical steroid receptor binding site is a DNA element with an AGGTCA consensus sequence that binds one or two receptor molecules as monomers or as homodimers and/or heterodimers (Tsai and O’Malley, 1994). The binding sites can be arranged as single half sites or as inverted, direct, or everted repeats. Although DNA binding by EcR-USP is highly sequence specific, a surprising variety of sequences have been identified as ecdysteroid response elements in D. melanogaster (see Chapter 3.5), suggesting that variability of EcREs provides yet another level of specificity in 20E gene regulation (Riddihough and Pelham, 1987; Cherbas et al., 1991; Antoniewski et al., 1994, 1995, 1996; D’Avino et al., 1995; Horner et al., 1995; Lehmann and Korge, 1995). The DNA binding properties of the AaEcRAaUSP heterodimer have been systematically analyzed with respect to the effects of nucleotide sequence, orientation, and spacing between half sites in natural D. melanogaster and synthetic EcREs (Wang et al., 1998). AaEcR-AaUSP exhibits a broad binding specificity, forming complexes with inverted repeats (IRs) and direct repeats (DRs) of the nuclear receptor response element half-site consensus sequence AGGTCA separated by spacers of varying length. A single nucleotide spacer was optimal for both imperfect (IRhsp-1) and perfect (IRper1) inverted repeats. Spacer length was less important in DRs of AGGTCA (DR-0 to DR-5). Although 4 bp was optimal, DR-3 and DR-5 bound AaEcR-AaUSP almost as efficiently as DR-4. Furthermore, AaEcRAaUSP also bound DRs separated by 11–13 nucleotide spacers. Cotransfection assays utilizing CV-1 cells have demonstrated that the mosquito EcRUSP heterodimer is capable of transactivating reporter constructs containing either IR-1 or DR-4.

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The levels of transactivation are correlated with the respective binding affinities of the response elements (IRper-1 > DR-4 > IRhsp-1). Taken together, these analyses predict broad variability in the EcREs of mosquito ecdysteroid responsive genes (Wang et al., 1998). These analyses enabled the identification of a functionally active EcRE that binds the heterodimer EcR-USP in the A. aegypti Vg gene. A direct repeat with a 1 bp spacer (DR-1) with the sequence AGGCCAaTGGTCG is the major part of the EcRE in the Vg gene (Martin et al., 2001a). 3.9.2.4.5. Ecdysteroid early response genes The A. aegypti E75 homolog consists of three overlapping transcription units encoding E75A, E75B, and E75C isoforms (Pierceall et al., 1999), which are members of the nuclear receptor family. As in D. melanogaster, these E75 transcripts arise by alternative promoter usage and splicing of transcription unit-specific 50 exons onto a set of shared downstream exons. All three transcripts are induced at the onset of vitellogenesis by a blood meal and are highly expressed in the mosquito ovary and fat body, which suggests that they are involved in the regulation of oogenesis and vitellogenesis, respectively. Furthermore, in vitro fat body experiments have demonstrated that AaE75 isoforms are induced by 20E. In contrast to the yolk protein transcripts, which show only a small gradual increase during the first 4 h PBM, E75 transcripts exhibit a sharp rise immediately after the onset of vitellogenesis, reaching a peak at 3–4 h PBM, when the hormone titer shows a first peak. After falling from their initial peak, the level of mosquito E75 transcripts gradually increase again, reaching the second peak at 18–24 h PBM, which coincides with the accumulation of yolk protein RNA in the fat body (Figure 9). E75 isoform transcription is appropriately 10 times more sensitive to 20E than is the Vg gene (Pierceall et al., 1999). Two isoforms of the homolog to the D. melanogaster transcription factor E74, which share a common C-terminal Ets DNA binding domain, yet have unique N-terminal sequences, are present in the mosquito. They exhibit a high level of similarity to DmE74 isoforms A and B and show structural features typical for members of the Ets transcription factor superfamily (Sun et al., 2002). Furthermore, both mosquito E74 isoforms bind to a D. melanogaster E74 binding site with the consensus motif C/ AGGAA. E74B mRNA reaches its peak at 24 h PBM and drops sharply thereafter, correlating with peak expression of the Vg gene. The AaE74B transcript is induced by a blood meal activated hormonal cascade in fat body and peaks at 24 h PBM, the peak of

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vitellogenesis (Figure 9). However, unlike E75 transcripts, E74B does not have an early peak at 3–4 h. AaE74A is activated at the termination of vitellogenesis and exhibits a peak at 36 h PBM in the fat body and 48 h PBM in the ovary. The AaE74A and AaE74B isoforms most likely play different roles in regulation of vitellogenesis in mosquitos, as an activator and a repressor of YPP gene expression, respectively. Both AaE74 isoform mRNAs are induced by 20E in fat body in vitro and display superinduction when cycloheximide is applied together with 20E (Sun et al., 2004). While both E74 isoforms are capable of binding to the E74 consensus sequence, only E74B induces a reporter gene expression through the consensus E74 binding site in cell transfection assays. Furthermore, a transient transfection assay using the Vg promoter region containing putative E74 binding sites has shown that E74B functions as an activator, whereas E74A serves as a repressor (Sun et al., 2004). The BR-C plays a key role in the genetic control of 20E responses. The BR-C encodes a family of DNA binding proteins that share a common (core) N-terminus fused by alternative splicing to one of four pairs of C2H2-type zinc finger domains, Z1, Z2, Z3, and Z4. All four BR-C isoforms are present in A. aegypti (Chen, Zhu, and Raikhel, unpublished data). In female fat body, expression of Z1, Z2, and Z4 is induced immediately after a blood meal. Both Z1 and Z4 transcripts exhibit a peak at 24 h PBM. The Z2 transcript exhibits a sharp increase after onset of vitellogenesis, reaching initial peak at 8 h PBM. After an initial decline, the level of Z2 rises again at 24 h PBM (Figure 9). The expression of BR-C Z1, Z2, and Z4 in the fat body is attenuated between 24 h and 36 h PBM. Z3 transcript expression in the fat body is very low throughout the vitellogenic cycle. The overall level of Z4 has been found to be much higher than those of the other isoforms. In the fat body in vitro experiments, the mRNA levels of all four isoforms were rapidly induced by 20E and increased in a dose dependent manner. Functional analysis using RNAi experiments has suggested that Z1 and Z4 serve as repressors and that Z2 serves as an activator of Vg gene expression. Remarkably, RNAi knockdown of Z1 or Z4 extends Vg expression to 36 h PBM, which suggests that these factors regulate hormone dependent timing of this gene transcription (Chen, Zhu, and Raikhel, unpublished data). Analyses of 20E sensitivity by transcripts of several key mediators and their expression profiles in vitellogenic fat body of the mosquito have revealed potential roles in the 20E gene regulatory hierarchy

cascade, thus providing a road map for future studies (Pierceall et al., 1999; Kapitskaya et al., 1998; Li et al., 2000; Wang et al., 2000b, 2002; Sun et al., 2002, 2004) (Figure 9). At the top of the 20E cascade, AaEcRB, the dominant isoform for initiating mosquito vitellogenesis in vivo, exhibits high sensitivity to 20E at 108 M, and it is expressed predominantly during the first hours after blood meal activation of vitellogenesis (Wang et al., 2002) (Figure 9). Its obligatory partner AaUSP-B, early gene products AaE74B and AaE75A, and BR-C isoforms constitute the group with the second highest sensitivity to 20E at 107 M (Pierceall et al., 1999; Wang et al., 2000; Sun et al., 2004; Chen, Zhu, and Raikhel, unpublished data). AaEcR-A, AaE74A, and the early late gene AaHR3 form the lowest 20E sensitivity group, being maximally activated at 106 M, which is similar to the target genes Vg, VCP, and VCB (Figure 9). 3.9.2.4.6. Hormonal enhancers in the regulatory region of mosquito yolk protein genes Transcriptional activation of hormonally controlled genes in specific tissues depends on interactions between sequence specific transcription factors and enhancer/promoter elements of these genes. Analysis of the 50 -upstream regulatory region of the mosquito Vg gene has revealed putative binding sites for EcR-USP and the early genes, E74, E75, and BR-C, which indicate this gene is regulated through a combination of direct and indirect hierarchies (Figure 10). Analyses of D. melanogaster and A. aegypti transformations, as well as DNA binding assays, have identified cis-regulatory sites in the Vg gene for stage and fat body specific activation via a blood meal triggered cascade (Kokoza et al., 2001). Three regulatory regions in the 2.1 kb, 50 region of the Vg gene are required for its blood meal activation and high-level expression (Figure 11). The proximal region, adjacent to the basal transcription start site, contains binding sites for several transcription factors that direct tissue and stage specific expression: EcR/USP, GATA transcription factor (GATA), CAAT binding protein (C/EBP), and hepatocyte nuclear factor 3/forkhead transcription factor (HNF3/ fkh). It appears that a combinatorial action of these transcription factors brings about fat body specific expression. EcR/USP acts as a timer, allowing the gene to be turned on, but the level of expression driven through this response element is low. The median region contains the sites for early gene factors E74 and E75, and transgenic studies have shown that this region is required for a stage specific hormonal enhancement of the Vg gene expression

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Figure 10 Direct and indirect regulation of vitellogenin (Vg) gene by 20-hydroxyecdysone in the mosquito fat body. The activated ecdysteroid receptor, consisting of EcR-B/USP-B binds to the EcRE response element in the regulatory region of the Vg gene. This binding is required to permit the Vg gene expression. After binding 20E, the EcR-USP heterodimer also activates early genes, E74, E75, and BR-C. The products of these genes E74-B, E75-A, and BR-C Z2 act as powerful activators of the Vg gene. (Based on Pierceall, W.E., Li, C., Biran, A., Miura, K., Raikhel, A.S., et al., 1999. E75 expression in mosquito ovary and fat body suggests reiterative use of ecdysone-regulated hierarchies in development and reproduction. Mol. Cell. Endocrinol. 150, 73–89; Kokoza, V.A., Martin, D., Mienaltowski, M.J., Ahmed, A., Morton, C.M., et al., 2001. Transcriptional regulation of the mosquito vitellogenin gene via a blood meal-triggered cascade. Gene 274, 47–65; Martin, D., Wang, S.F., Raikhel, A.S., 2001b. The vitellogenin gene of the mosquito Aedes aegypti is a direct target of ecdysteroid receptor. Mol. Cell. Endocrinol. 173, 75–86; Sun, G.Q., Zhu, J.S., Li, C., Tu, Z.J., Raikhel, A.S., 2002. Two isoforms of the early E74 gene, an Ets transcription factor homologue, are implicated in the ecdysteroid hierarchy governing vitellogenesis of the mosquito, Aedes aegypti. Mol. Cell. Endocrinol. 190, 147–157; L. Chen, J. Zhu, and A.S. Raikhel, unpublished data.)

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(Kokoza et al., 2001). Furthermore, in vitro cell transfection experiments have demonstrated that the ecdysteroid receptor and E74B act synergistically to bring about a high-level, 20E activated Vg gene expression (Sun and Raikhel, unpublished data). Finally, the distal portion is characterized by multiple response elements for a GATA transcription factor (Martin et al., 2001b). In transgenic experiments using both A. aegypti and D. melanoagaster, this GATA- rich region is required for high expression levels characteristic to the Vg gene (Figure 11). Recent analyses of the A. aegypti Vg promoter region have identified many binding sites for the BR-C transcription factors: one for Z1, two for Z2, three for Z3, and seven for Z4. When BR-C isoform expression vectors were introduced into Kc cells together with EcR and USP expression vectors in the presence of 20E, Z1 and Z4 repressed the activation mediated by EcR-USP, while Z2 enhanced this activation. The role of Z3 was negligible. When the Z1 binding site was mutated, luciferase activity driven by the Vg promoter was no longer inhibited by the overexpression of Z1. After removal of Z2 binding sites in the Vg promoter, Z2 lost the capability to increase luciferase activity. However, because there are seven binding sites of BR-C Z4 on the Vg promoter, the repression of Vg promoter by Z4 could not be completely released after mutation of all the Z4 binding sites. These experiments support the RNAi test performed in vivo and suggest that BR-C Z1 and Z4 serve as repressors while Z2 is an activator of the Vg gene expression. Z4 may not directly bind to the Vg promoter (Chen, Zhu, and Raikhel, unpublished data).

Figure 11 Schematic illustration of the regulatory regions of the Aedes aegypti Vg gene. Numbers refer to nucleotide positions relative to the transcription start site. Binding sites for hormonal transcription factors are depicted in red and those for tissue specific factors in green. C/EBP, response element of C/EBP transcription factor; EcRE, ecdysteroid response element; E74 and E75, response elements for respective early gene product of the ecdysone hierarchy; GATA, response element for GATA transcription factor; HNF3/fkh, response element for HNF3/forkhead factor; Vg (purple), coding region of the Vg gene; Z1, Z2, and Z4 are binding sites for corresponding Broad-Complex isoforms. (Modified with permission from Kokoza, V.A., Martin, D., Mienaltowski, M.J., Ahmed, A., Morton, C.M., et al., 2001. Transcriptional regulation of the mosquito vitellogenin gene via a blood meal-triggered cascade. Gene 274, 47–65.)

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3.9.2.4.7. Competence to 20E response Female mosquitos, such as A. aegypti, require a previtellogenic preparatory period to attain the capability for host seeking behavior and blood feeding. During this time, the fat body becomes competent for massive yolk protein synthesis and secretion, and the ovary for accumulation of YPs. In the fat body, this process is manifested in the development of the endoplasmic reticulum and Golgi complexes, proliferation of ribosomes, and increase in cell ploidy (Dittmann et al., 1989; Raikhel and Lea, 1990). JH III titers rise tenfold over the first 2 days after emergence and then slowly decline over the next 5 days. Blood ingestion causes an immediate decline of JH, which falls to its lowest level at 24 h PBM (Shapiro et al., 1986). Activation of fat body nucleoli for ribosomal RNA production and ribosomal production is blocked by removal of the CA in newly eclosed adult females, but it can be restored by either implantation of CA or topical application of JH III to allatectomized females. These developmental events most likely are controlled by JH from the CA (Raikhel and Lea, 1990, 1991). The exposure of a newly emerged female mosquito to JH III is essential for the fat body to become responsive to 20E (Flanagan et al., 1977; Li et al., 2000; Zhu et al., 2003a). To dissect the molecular mechanism governing the acquisition of competence for vitellogenesis in the A. aegypti fat body, the mosquito homolog of the ecdysteroid response competence factor, bFTZ-F1, in D. melanogaster has been cloned (Li et al., 2000). During metamorphosis in D. melanogaster, the stage specificity of the genetic response to 20E is set up by bFTZ-F1, an orphan nuclear receptor. Ectopic bFTZ-F1 expression leads to elevated transcription levels for 20E-inducible BR-C and the early genes E74 and E75. bFTZ-F1 mutants pupate normally in response to the late larval 20E pulse but display defects in stage specific responses to the subsequent 20E pulse in prepupae (Lam et al., 1997; White et al., 1997). Mosquito bFTZ-F1 is transcribed highly in the late pupa and in the adult female fat body during pre- and postvitellogenic periods, when ecdysteroid titers are low, and the transcripts nearly disappear in midvitellogenesis, when ecdysteroid titers are high. Each rise in the level of bFTZ-F1 transcripts is preceded by a high expression of another nuclear receptor (HR3) that coincides with the 20E peaks (Kapitskaya et al., 2000; Li et al., 2000). This observation is consistent with the role of HR3 in D. melanogaster, which facilitates induction of bFTZ-F1 in mid-prepupa. Although the genetic tools are limited for the A. aegypti, experiments with the fat body in vitro

lend support at the functional level to the hypothesis that FTZ-F1 serves as a competence factor for the ecdysteroid response initiating vitellogenesis. In these experiments, FTZ-F1 transcription is inhibited by 20E and is superactivated by its withdrawal (Li et al., 2000). Electrophoretic mobility-shift assay analysis of nuclear extracts from A. aegypti fat body demonstrated that the onset of ecdysone response competence in this tissue is correlated with the appearance of the functional FTZ-F1 protein at the end of previtellogenic development (Li et al., 2000). Western blot analysis using antibodies to A. aegypti bFTZ-F1 show that the bFTZ-F1 protein is not detectable in the fat body nuclear extracts of newly emerged and 1-day-old mosquitos but is abundant at 3–5 days posteclosion (PE), and then not dectected shortly after blood feeding (Zhu et al., 2003a). Furthermore, when fat body from females was isolated at 6 h PE and incubated for 18 h in medium with JH III, the nuclei contained bFTZ-F1 protein, in contrast to controls incubated with acetone (Zhu et al., 2003a). Taken together, these findings indicate that JH III is quite likely the crucial factor modulating production of bFTZ-F1 protein in the fat body through posttranscriptional regulation of its gene in the previtellogenic stage (Figure 12). To confirm that bFTZ-F1 may encode a competence factor in the fat body of female A. aegypti, it was silenced by RNAi, which involves the introduction of homologous double-stranded RNA (dsRNA) in order to target specific mRNAs for degradation (Zhu et al., 2003a). Compared with naı¨ve females, the mRNA and protein levels of bFTZ-F1 declined substantially in females treated with bFTZ-F1 dsRNA but not in those treated with control dsRNA, which indicates that bFTZ-F1 was selectively inhibited by RNAi. Expression of the Vg gene was dramatically diminished after blood feeding in the bFTZ-F1 dsRNA treated mosquitos. These results suggest that A. aegypti bFTZ-F1 is essential for the stage specific 20E response in fat body during vitellogenesis, which is reminiscent of the role bFTZ-F1 plays in D. melanogaster during the prepupa to pupa transition. Functional analysis of bFTZ-F1 by the RNAi technique suggests that bFTZ-F1 is in fact a competence factor that defines the stage specific 20E response during vitellogenesis in the mosquito (Zhu et al., 2003a). Despite extensive studies, the mode of JH action is still not well understood. During previtellogenic development in female mosquitos, JH controls the synthesis of bFTZ-F1 protein, but this posttranscriptional control of gene expression can occur at one or more levels: pre-mRNA splicing pattern, mRNA

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Figure 12 bFTZ-F1, the orphan nuclear factor (see Chapter 3.5), is implicated as a competence factor for stage specific response to ecdysteroid during vitellogenesis in the Aedes aegypti fat body. The AaFTZ-F1 mRNA is present at late pupal and previtellogenic stages in newly eclosed females; however, the appearance of active AaFTZ-F1 factor coincides with the onset of competence for 20E response (Li et al., 2000; Zhu et al., 2003a). (Reprinted with permission from Raikhel, A.S., Kokoza, V.A., Zhu, J.S., Martin, D., Wang, S.F., et al., 2002. Molecular biology of mosquito vitellogenesis: from basic studies to genetic engineering of antipathogen immunity. Insect Biochem. Mol. Biol. 32, 1275–1286; ß Elsevier.)

stability, mRNA transport, or translation rate. Although the mechanism underlying this regulation remains obscure, these findings represent another step towards understanding the mode of JH action. 3.9.2.4.8. Regulation of vitellogenesis cyclicity by ecdysteroid mediated signaling through heterodimerization with the RXR homolog Ultraspiracle In many insects, vitellogenesis and egg maturation arecyclic and, as a consequence, these insects produce batches of eggs, unlike others that lay eggs continuously. Generally, the mode of egg production – continuous or cyclic – depends upon feeding and lifestyle adaptations. Cyclic egg production occurs in all insect genera irrespective of the type of hormonal control. In insects, in which reproduction is governed by JH, these mechanisms are poorly understood at the molecular level. However, for insects with cyclic egg production that depends primarily on ecdysteroids, recent studies of the mosquito, A. aegypti, have shed some light on these mechanisms. Zhu et al. (2000, 2003b) and Miura et al. (2002) have demonstrated that during mosquito vitellogenesis, two nuclear receptors, HR38 and Svp, regulate the cyclicity of ecdysteroid mediated signaling via heterodimerization with USP. Both AaEcR and AaUSP proteins are abundant in nuclei of the

previtellogenic female fat body at the state of arrest; however, the EcR-USP heterodimer capable of binding to the specific ecdysone response elements is barely detectable in these nuclei. Studies have shown that EcR is a primary target of 20E signaling modulation in target tissues at the state of arrest. A possible mechanism through which the formation of ecdysteroid receptor activity can be regulated is a competitive binding of other factors to either EcR or USP. Indeed, at this stage, AaUSP exists as a heterodimer with the orphan nuclear receptor, AHR38. This protein is a homolog of the D. melanogaster DHR38 and vertebrate NGFI-B/Nurr1 orphan receptors, and it acts as a repressor by disrupting EcR binding to DNA response elements and by interacting strongly with AaUSP. However, in the presence of 106 M of 20E, EcR efficiently displaces AHR38 and forms an active heterodimer with USP, as happens after a blood meal (Figures 13 and 14). To regulate cyclicity in egg production, termination of Vg gene expression is needed in female mosquitos, so that egg maturation and deposition can be completed and the arrest stage restored until another blood meal can be obtained. To identify other negative regulators of the AaEcR/AaUSP mediated 20E response during vitellogenesis, a

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mosquito homolog (AaSvp) of the vertebrate orphan nuclear receptor, chicken ovalbumin upstream promoter transcription factor (COUP-TF), and D. melanogaster Seven-up (Svp) has been cloned (Miura et al., 2002). Coimmunoprecipitation experiments with nuclear extracts (Zhu et al., 2003b) have clearly shown that in the female fat body AaSvp associates only with AaUSP. Furthermore, formation of AaSvp-AaUSP heterodimers occurs in a precise, timely manner at 30–33 h PBM, after the 20E titer declines to the previtellogenic level and when the expression of YPP genes, including Vg, is terminated (Figure 13). In vitro experiments have suggested that the declining titer of 20E could be a critical factor facilitating the formation of Svp-USP heterodimers (Zhu et al., 2003a). Mosquito Svp, thus, represses the 20E mediated transactivation of vitellogenesis through its heterodimerization with USP (Zhu et al., 2003b). The modulation of vitellogenesis cyclicity appears to be regulated through alternative heterodimerization of the RXR homolog USP that blocks ecdysteroid mediated signaling (Figure 14). AaUSP exerts its functions by associating with distinct partners at different stages of vitellogenesis. During the arrest stage, AHR38 prevents the formation of the functional EcR complex by sequestering AaUSP,

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which blocks 20E dependent transactivation. After a blood meal, the AaEcR-AaUSP heterodimerization becomes dominant, and AaEcR-AaUSP binding to EcREs on the Vg gene permits its expression. When vitellogenesis proceeds to the termination stage, falling 20E titers shift AaUSP heterodimerization towards AaSvp, repressing USP-based hormone responses (Figure 14). In summary, these studies clearly show that the cyclicity of vitellogenesis in the mosquito fat body is regulated through USP, which sequentially forms inactive or active heterodimers with either repressors (HR38 and Svp) or the activator (EcR) and directly affects ecdysteroid mediated signaling. 3.9.2.5. Molecular Endocrinology of Vitellogenesis in the Cyclorrhaphan Diptera

3.9.2.5.1. Vitellogenesis and hormones in D. melanogaster Like the mosquito, the fat body in fruit flies matures and differentiates at eclosion under the control of JH, but it immediately begins to synthesize YPs and secrete them into the hemolymph. Simultaneously, the uptake of YPs is initiated by developing oocytes that mature into eggs, which are fertilized and laid continuously. If the fly does not mate or has enough food, mature eggs are retained, and oocytes

Figure 13 Aedes aegypti Ultraspiracle (AaUSP) interacts sequentially with a repressor AHR38, an activator AaEcR, and then with repressor AaSvp in the fat body during the vitellogenic cycle. Nuclear extracts were prepared from the fat bodies of 500 adult females for each time point. (a) Coimmunoprecipitation analysis. A protein aliquot equivalent to 100 mosquitos was incubated with anti-Drosophila USP monoclonal antibody. The resulting immune complexes were then precipitated by the addition of protein A-agarose beads. After extensive washing, immune complexes were dissociated and separated by means of SDS-PAGE followed by immunoblotting using rabbit anti-AHR38, rabbit anti-AaEcR, chicken anti-AaSvp antibodies, or the respective preimmune sera. In vitro translated proteins (AHR38, AaEcR-B, and AaSvp) were used as controls in Western blot analysis. (b) Western blot analysis of USP proteins during the vitellogenic cycle shows that two isoforms of USP are present throughout the vitellogenic cycle in fat body nuclei. A protein aliquot equivalent to 25 fat bodies was loaded in each lane for SDS-PAGE. Western blot analysis was performed using the anti-DmUSP monoclonal antibody. In vitro TNT expressed USP-A and USP-B proteins were used as positive controls. (Reproduced with permission from Zhu, J.S., Miura, K., Chen, L., Raikhel, A.S., 2003b. Cyclicity of mosquito vitellogenic ecdysteroid-mediated signaling is modulated by alternative dimerization of the RXR homologue Ultraspiracle. Proc. Natl Acad. Sci. USA 100, 544–549.)

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Figure 14 Heterodimerization of Ultraspiracle (USP), the obligatory partner of EcR, with repressor nuclear receptors during the first vitellogenic cycle in the mosquito Aedes aegypti. At the state of arrest, AaUSP was associated with AHR38, preventing activation of YPP production prior to blood feeding. After a blood meal, AaUSP heterodimerizes with AaEcR, and induces expression of YPP genes in the presence of an elevated 20E titer. Around 30–36 h PBM, AaSvp attaches to AaUSP, decelerating the massive protein synthesis. Top panel: stages of mosquito vitellogenic cycle; bottom panel: titers of juvenile hormone and ecdysteroids. (Modified with permission from Zhu, J.S., Miura, K., Chen, L., Raikhel, A.S., 2003b. Cyclicity of mosquito vitellogenic ecdysteroid-mediated signaling is modulated by alternative dimerization of the RXR homologue Ultraspiracle. Proc. Natl Acad. Sci. USA 100, 544–549.)

arrest at a previtellogenic stage and do not enter vitellogenesis (Bownes, personal communication). Despite the fact that in most cyclorrhaphan Diptera, including D. melanogaster, YPs are made in fat body and follicular epithelial cells surrounding the oocyte, the regulation of yp gene expression is different in these tissues (Bownes, personal communication). Ecdysteroids stimulate YP synthesis in the fat body of males and upregulate it in the fat body of females (Postlethwait et al., 1980; Jowett and Postlethwait, 1980; Bownes, 1982; Bownes et al., 1983). This is consistent with the finding that EcREs have been mapped to regions flanking the yp genes (Bownes et al., 1996). JH also increases YP synthesis in the fat body of females (Bownes and Blair, 1986; Bownes et al., 1987; Soller et al., 1997), but it does not induce YP synthesis in males. No potential JH elements have been found in these genes. Walker et al. (1991) have shown that ecdysteroids increased CATexpression of YP promoter CAT fusion in the S3 cell line. JH had no effect on the same reporter construct. Thus, it is unlikely that JH plays any role in directly affecting transcription of yp genes in the fat body. JH may play a role in the regulation of yolk protein synthesis by follicle cells, as shown in studies of the D. melanogaster mutant ap56f (Richard et al., 1998, 2001). Female ap56f produces

low levels of JH and normal to elevated levels of ovarian ecdysteroids and are fertile but have delayed vitellogenesis. JH application to female ap56f reversed the delay in vitellogenesis. Many of the response genes in the ecdysteroid regulatory hierarchy, including EcR/USP E74, E75, and BR-C, that have been shown to be crucial for 20E regulation of Vg gene expression in the fat body of A. aegypti have not been studied in D. melanogaster with respect to YP synthesis in the fat body. This is mostly because in normal, mated, wellfed females the genes are constitutively active rather than regulated in response to a blood meal (Raikhel et al., 2003; Bownes, personal communication). 3.9.2.5.2. Regulatory regions of yolk protein genes in D. melanogaster Relatively little is known about the cis-acting regions of the yp genes in any species of cyclorrhaphan Diptera. Several EcREs have been identified in the 50 and 30 regions and within the coding sequences of the yp genes of D. melanogaster (see Figures 15 and 16), and they appear to confer ecdysone inducible expression in males, when injected with 20E. These putative EcRE sequences are similar to those shown to bind the EcRE/USP heterodimer and lead to transcription of other genes in insect metamorphosis and the Vg

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Figure 15 Drosophila melanogaster yp genes 1 and 2 are located in close proximity in opposite orientation and share a regulatory region called the intergenic enhancer region. The 1225 bp intergenic enhancer region of the yp1 and yp2 genes contains four enhancer regions, the fat body enhancer (FBE), the hermaphrodite response region (HRR), and two ovary enhancers (OE1 and OE2); aef-1, the adult enhancer factor-1; bzip3; C/EBP, CAAT enhancer binding protein; dsx, doublesex binding site; EcRE, ecdysone response element; her, binding site for the hermaphrodite protein. (Data kindly provided by M. Bownes.)

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Figure 16 The Drosophila yp3 gene is separated from two others on the X-chromosome and has a separate regulatory region, the upstream enhancer 3. aef-1, adult enhancer factor 1; bbf, binding site for the box-B binding factor; EcRE, ecdysone response element. (Data kindly provided by M. Bownes.)

genes in the mosquito (Bownes et al., 1996). Although these EcREs have been mapped on the yp genes, it is not clear how much of the 20E inducibility of yp gene expression is a direct action through the binding of EcR and how much is due to downstream genes such as BR-C, E74, and E75 (Bownes, personal communication). Despite the fact that the hormonal and molecular control of yp gene transcription in D. melanogaster is poorly understood, the sex specific expression of these genes has been elucidated in detail (see Chapters 1.5 and 1.7). Doublesex (dsx) exerts the most important control of yp gene expression (Bownes and Nothiger, 1981; Coschigano and Wensink, 1993; Belote et al., 1985). Alternate transcript splicing of dsx produces different proteins in male and female adults, DsxM and DsxF, respectively, that have similar DNA binding domains but different C-terminal extensions (Baker and Wolfner, 1988; Burtis and Baker, 1989; MacDougall et al., 1995). The male and female proteins are essential for maintaining yp transcription in the female fat body and repressing it in males (Bownes and

Nothiger, 1981; Belote et al., 1985; Burtis et al., 1991; Bownes, personal communication). In males, 20E injection overrides sex determination inhibition as shown by the transient expression of yp genes in the fat body of males (Bownes et al., 1996; Bownes, personal communication). In females, yp gene expression in the ovarian follicle cells does not depend upon the sex determination pathway (Logan et al., 1989; Logan and Wensink, 1990; Lossky and Wensink, 1995), and 20E is probably a key hormone regulating this expression in such cells (Bownes, 2004). In D. melanogaster, yp genes 1 and 2 are located in close proximity in opposite orientation and share a regulatory region called the intergenic enhancer region (Figure 15). In females, expression of yp1 and yp2 occurs in fat body and follicle cells of developing egg chambers at stages 8–10 (Bownes, 2004). The 1225 bp intergenic enhancer region of these yp genes contains four enhancer regions: fat body enhancer (FBE), hermaphrodite response region (HRR), and two ovary enhancers (OE1 and OE2) (Logan et al., 1989; Logan and Wensink,

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1990; Garabedian et al., 1985, 1986; Abrahamsen et al., 1993; Lossky and Wensink, 1995). FBE drives expression of these genes in fat body, and HRR responds to the hermaphrodite gene that acts along with Dsx to control their sex specific expression (Li and Baker, 1998). OE1 and OE2 direct expression of these genes in follicle cells (Logan et al., 1989; Logan and Wensink, 1990; Lossky and Wensink, 1995). With base pairing in respect to the yp1 start site, several binding sites for activator or repressor molecules have been mapped, and binding has been demonstrated for some of these molecules (Figure 16). A putative binding site for bzip3 overlaps a weak binding site for a Dsx, dsxC. Two overlapping binding sites for the mammalian CAAT enhancer binding protein (C/EBP) and the adult enhancer factor-1 (AEF-1) have been found in other fat body enhancers and the Adh gene of the mammalian liver to activate or repress transcription, respectively (Abel et al., 1992). These two sites, in turn, overlap the strongest binding site for Dsx, dsxA (Bownes et al., 1996). Further upstream, between 322 and 1225 bp, up to the yp2 transcription start site, lies the HRR, which contains the binding site for the hermaphrodite protein, and OE1. A putative EcRE lies within OE1. A second EcRE is found between 482 and 494 bp of the HRR region. OE2 is located at the start site for yp2 transcription (Logan et al., 1989; Logan and Wensink, 1990; Lossky and Wensink, 1995; Bownes, ; Bownes, personal communication). The D. melanogaster yp3 gene is separated from two others on the X-chromosome and has a separate regulatory region, the upstream Enhancer 3 (Hutson and Bownes, 2003). The organization of the regulatory regions of this gene differs from those of yp1 and yp2, resembling instead those in Musca and Calliphora yp genes (Hutson and Bownes, 2003). The 703 bp upstream enhancer sequence of the yp3 transcription start site contains three regions that confer tissue specificity, sex specificity, and ovary specificity (Figure 16). The 419 bp fat body enhancer 3 (FBE3) is the furthest 50 regulatory region spanning from 704 to 285 bp and controls expression of yp3 in the female fat body (Hutson and Bownes, 2003). Most 50 in the FBE3 is the binding site for dsxA. Downstream of the dsxA site is the overlapping binding site, also found in FBE1/2, for the activator molecule CAAT enhancer binding protein (C/EBP), and the repressor, adult enhancer factor 1 – AEF-1 (Falb and Maniatis, 1992a, 1992b). The region between 498 and 252 bp is essential for sex specificity. There are two binding sites for the box-B binding factor (BBF), a transcriptional activator (Abel et al., 1992). A possible

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mechanism also involves a putative homolog of the unknown activator R that has also been found to be the final determinant needed for female specific fat body expression of yp1 and 2 (Ronaldson and Bownes, 1995; Hutson and Bownes, 2003). Hutson and Bownes (2003) have analyzed the regulation of yp3 gene expression. JH increased stability of yp3 transcripts, and 20E plays an important role in regulating transcription. Three putative EcREs have been mapped in FBE3 and yp3 (Hutson and Bownes, 2003). Immediately 30 to the FBE3 is the ovarian enhancer 3 (OE3) that is necessary for regulation of yp3 transcription in the follicle cells of the developing egg chamber between stages 8 and 10. Within this region is an EcRE, and a second EcRE is located 30 of the intronic sequence of yp3. The final EcRE is found within the second exon of yp3. 20E can override the negative effects of DsxM while maintaining the tissue specificity of yp3 gene expression (Hutson and Bownes, 2003). Experiments using upstream, downstream, and coding sequences of yp3 fused to reporter genes agree well with the locations of the putative EcREs, because there are ecdysteroid inducible sites upstream, downstream, and in the coding region (Hutson and Bownes, 2003). 3.9.2.6. Hormones and Ovarian Maturation

3.9.2.6.1. Previtellogenic development of the follicle Several aspects of ovarian previtellogenic development in insects are controlled by JH. Stimulation of previtellogenic growth of follicles by JH is best supported by past studies (Strong, 1965; Gwardz and Spielman, 1973; Tobe and Pratt, 1975; Tobe and Stay, 1977; Lanzrein et al., 1978; Moobola and Cupp, 1978; Tobe and Langley, 1978; McCaffery and McCaffery, 1983). Differentiation of the follicular epithelium is another aspect of oocyte development that is regulated by JH (Davey and Hubner, 1974; Abu-Hakima and Davey, 1975; Elliot and Gillot, 1976; Koeppe and Wellman, 1980; Koeppe et al., 1980). In M. domestica, JH is released soon after adult emergence, and it is essential for ovarian maturation, since without it ovaries remain immature (Adams, 1974, 1980). JH stimulates endopolyploidy in the nurse cells, primes the oocyte for growth, and potentially plays a role in follicle morphogenesis. Although, JH affects the previtellogenic stages of M. domestica oogenesis, it does not induce vitellogenesis. JH also is required for oocyte development in the mosquito A. aegypti, where it controls growth and differentiation of the follicular epithelium. These events are blocked by the removal of CA and restored by either implantation of CA or application of JH (Raikhel and Lea, 1991).

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3.9.2.6.2. The role of 20E in ovarian development and egg maturation in Diptera Ecdysteroids are produced by the ovaries of many insects, including mosquitos and flies (Hagedorn et al., 1975; Bownes et al., 1984; Hagedorn, 1985). However, their role in ovarian development is not completely understood. In D. melanogaster, ecdysteroids can have an antagonistic effect on oocyte progression and lead to apoptosis of egg chambers. Ecdysteroid mutants, such as ecd1 (Redfern and Bownes, 1982) and l(3)3DTS (Walker et al., 1987), have defects in the progress of oogenesis. One of the early genes in the ecdysteroid regulatory hierarchy, BR-C, is crucial at several stages of oogenesis. It is first expressed in all follicle cells surrounding the oocyte prior to the control point (Deng and Bownes, 1997), and this early BR-C induction may be controlled by ecdysteroids. An exciting development is the isolation of the D. melanogaster dare gene that encodes adrenodoxin reductase, which is expressed in the ovary and plays a key role in ecdysteroid biosynthesis (Freeman et al., 1999) (see Chapter 3.3). These authors postulated that ovarian expression of dare provides a maternal source of this enzyme to the developing embryo. Since it is first expressed just prior to activation of the BR-C, it is also possible that it could also support the synthesis of ecdysteroids, which would activate BR-C in nearby follicle cells at this stage of oogenesis. This scenario suggests the existence of an egg chamber autonomous timing event that regulates when and where ecdysteroids are available to control oocyte progression. BR-C expression in oogenesis is crucial for the regulation of endoreplication of polyploid follicle cells, specific amplification of the chorion

genes, and later for positioning of the chorionic appendages (Deng and Bownes, 1997; Tzolovsky et al., 1999). Although BR-C is regulated by ecdysteroids in early oogenesis, the epidermal growth factor signaling pathway in the egg chamber is crucial for positioning the late BR-C expression pattern, which in turn positions the chorionic appendages (Deng et al., 1999). Since the effects of 20E are mediated by EcR and USP, mutations in these genes in D. melanogaster affect oogenesis. EcR-A and Ec-R-B1 are expressed in the nurse cells and follicle cells throughout oogenesis (Deng, Mauchline, and Bownes, personal communication). A temperature sensitive mutant affecting these isoforms, EcRA483T, reduced egg laying, induced abnormal egg clusters, and led to a loss of vitellogenic stages (Carney and Bender, 2000). Loss of EcR in germline clones leads to egg chamber arrest at around stage 6–7 of follicle development (previtellogenic stage). Carney and Bender (2000) proposed the existence of an ecdysone dependent checkpoint at this stage. Buszczak et al. (1999) have shown that the early ecdysone response genes, E75 and E74, along with the BR-C, are crucial for egg chamber morphogenesis in mid-oogenesis. Germline clones of cells lacking E75, dare, or EcR functions results in the degeneration of these egg chambers at stage 8 (Buszczak et al., 1999). 3.9.2.6.3. Regulation of patency by follicle cells The appearance of large intercellular spaces in the follicular epithelium during vitellogenesis, a phenomenon termed patency, has been reported in many insects, from the less modified species having panoistic ovaries, like B. germanica (Figure 17)

Figure 17 Patency of follicle cells in Blattella germanica during yolk protein uptake. (a, b) Sections at the equatorial zone in basal oocytes of (a) 0.68 mm length (3 days old) and (b) 1.52 mm length (5 days old). (c) Relation between patency index (PI) and basal oocyte length. PI ¼ SE/SC, where SE is the surface of the intercellular spaces stained with Evans’ blue, and SC the surface of the follicle cells. (Reprinted with permission from Pascual, N., Cerda`, X., Benito, B., Toma´s, J., Piulachs, M.D., et al., 1992. Ovarian ecdysteroid levels and basal oocyte development during maturation in the cockroach Blattella germanica (L.). J. Insect Physiol. 38, 339–348, with permission from Elsevier.)

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(Pascual et al., 1992) to the most modified ones having polytrophic ovaries, like flies or mosquitos (Wyatt and Davey, 1996). Studies on Rhodnius prolixus by Davey and co-workers have suggested that patency is under JH control (Wyatt and Davey, 1996). Further studies in this bug have shown that JH increases the activity of Naþ/Kþ ATPase in membrane preparations from vitellogenic follicle cells incubated in vitro, and experiments with inhibitors and activators of protein kinase C indicated that this enzyme is involved in the activation of the ATPase (Sevala and Davey, 1989). Incubation of membrane preparations with JH I resulted in the phosphorylation of a 100 kDa protein, a process that was dependent on protein kinase C and was inhibited by ouabain. Based on these experiments, it was suggested that the 100 kDa protein could be the asubunit of the ATPase (Sevala and Davey, 1993). According to this model, JH would act on the follicle cells via a protein kinase C dependent cascade to stimulate a membrane bound Naþ/Kþ ATPase (Figure 18). Then, changes in the ionic balance would cause water loss and shrinkage of the cells with the associated apparition of large intercellular spaces. An equivalent system seems to operate in the locust L. migratoria (Davey et al., 1993) and the mealworm beetle, Tenebrio molitor (Webb et al., 1997). The model proposed to explain these mechanisms acting in follicle cell patency (Figure 18) suggests the existence of a membrane receptor that binds JH. The first efforts to characterize such a receptor were carried out in R. prolixus. In this species, Ilenchuk and Davey (1985) have shown that JH I binds to membrane preparations of follicle cells with a Kd of 6.54 nM in a

Figure 18 Model for the action of JH on the follicle cell membrane. The binding of JH to the JH receptor (JHR) on the outer surface of the membrane triggers a cascade involving a G protein, with which the receptor site is closely associated. Protein kinase C (pkC) is activated through phosphodiesterase (PDE) and diacylglycerol (DAG) and phosphorylates the a-subunit of the NaþKþ ATPase, thus activating the enzyme. (Modified from Wyatt, G.R., Davey K.G., 1996. Cellular and molecular actions of juvenile hormone. II. Roles of juvenile hormone in adult insects. Adv. Insect Physiol. 26, 1–155.)

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specific and saturable fashion. Neither JH II nor JH III exhibited biological activity, and they did not compete for the binding site (Ilenchuk and Davey, 1987). A 35 kDa protein in solubilized follicle cell membranes from L. migratoria was found to bind a photoaffinity analog of JH III, [3H]EFDA (Sevala et al., 1995). The membrane preparations bound [3H]JH III with a Kd of 3.68 nM in a specific and saturable fashion, and the binding of EFDA was blocked when membranes were previously treated with JH III. Interestingly, EBDA, a photoaffinity analog of JH I, did not show any specific binding in equivalent assays; EHDA, a photoaffinity analog of JH II, bound specifically to a 35 kDa protein, and MDK, a photoaffinity analog of methoprene, bound specifically to a different 17 kDa protein (Sevala et al., 1995). Although relatively small, these JH binding proteins may serve as membrane receptors; certainly the 35 kDa protein is within the size range of G protein-coupled receptors (see Chapter 5.5). Further investigations are required to elucidate the nature of these proteins. Webb and Hurd (1995) have reported specific and saturable binding of JH III to microsomal preparations of vitellogenic follicle cells from the beetle, T. molitor. Results of Scatchard analysis indicated the occurrence of two sites of different affinity, one with a Kd of 10 nM and the other one of 400 nM. Both sites showed modest affinity for JH I. The specificity of JH binding by follicle cell membranes from the above species is intriguing, given the structural similarity of JH homologs: R. prolixus to JH I only, L. migratoria to JH III and II, and T. molitor to JH III and also JH I. Coincidently, JH III is found in L. migratoria and T. molitor, while that of R. prolixus is unknown. 3.9.2.6.4. Development of oocyte endocytic complex and endocytosis When YPs reach the plasma membrane of oocytes (oolemma), they are internalized into the ooplasm through receptor mediated endocytosis, via coated pits and vesicles, and transferred to early endosomes in the cortical ooplasm. From early endosomes they are conveyed to yolk spheres, while receptors are recycled back to the oocyte surface (Raikhel and Dhadialla, 1992; Sappington and Raikhel, 1998a; Snigirevskaya and Raikhel, 2004; Raikhel et al., 2002). In D. melanogaster, the endocytic pathway leading to the production of yolk spheres was visualized by Giorgi et al. (1993) with exposure to peroxidase in vivo or in vitro. The Golgi apparatus and the yolk spheres were then labeled by fixation with osmium zinc iodide (OZI). Starvation resulted in the OZI labeling being restricted to the Golgi apparatus and to an

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extended tubular network, whereas feeding or treatment with JH caused the yolk spheres to become labeled with OZI and to incorporate peroxidase. Similar results were obtained in equivalent experiments with the mutant ap4, which is defective in JH biosynthesis, or with vitellogenic follicular tissue incubated in vitro and treated or not with JH. The results of this study indicated that JH facilitates endocytic uptake by inducing the fusion of coated vesicles and tubules with the yolk spheres (Giorgi et al., 1993). D. melanogaster ap4 mutant flies produce low levels of JH and ovarian ecdysteroids and are infertile (Richard et al., 1998, 2001) This situation suggests that JH has a prominent role in regulating YP uptake by the ovary, given that the endocytic organelles are absent in ap4 oocytes and YPs are present at normal levels in the hemolymph. The development of endocytic organelles can be restored in ap4 flies by application of JH in a fashion similar to that of mosquito. In insects with continuous egg production such as D. melanogaster, it is not clear whether JH regulates the formation of endocytic organelles, stimulates the endocytosis itself or acts in both events, because these events are difficult to separate. In insects with cyclic egg production, early developmental and vitellogenic events in egg chambers are separated at the arrest stage, which is broken by either food intake or other stimuli. Normally, these insects mature only a single egg per ovariole in each cycle. Raikhel and Lea (1985) have shown that during the JH dependent previtellogenic period of follicle development in the mosquito A. aegypti, a highly specialized endocytic complex consisting of microvilli, numerous coated vesicles, and endosomes appears in the oocyte cortex. The formation of this endocytic complex is controlled by JH III. It was blocked by CA ablation at eclosion, but restored by either CA implantation or the application of JH III. Analysis of several genes involved in receptor mediated endocytosis of YPs, e.g., clathrin heavy chain (Kokoza and Raikhel, 1997), Vg receptor (Sappington et al., 1996), and ovarian lipophorin receptor (Cheon et al., 2001), has revealed that their expression occurs very early in the previtellogenic development of mosquito ovaries, even before the formation of the primary follicle. Immunocytochemistry, however, shows that the Vg receptor is present as the development of the endocytic complex occurs in the oocyte cortex (Sappington et al., 1995). These findings suggest that JH likely acts at the posttranscriptional level on these genes in a manner similar to that in the fat body (Zhu et al., 2003a).

3.9.2.6.5. The ovary as an endocrine organ Many different cell types and tissues make up the insect ovary, and any of these cells or tissues may produce factors that affect itself (autocrine), neighboring cells (paracrine), or more distant cells (endocrine). The ovary is a primary source of ecdysteroids in female insects, as demonstrated in many different insect groups (e.g., Whiting et al., 1997; Marti et al., 2003), and autocrine, paracrine, and endocrine effects have been reported or ascribed to ovarian ecdysteroids in female insects (Hagedorn, 1985; Lanot et al., 1989). The follicle cells surrounding the oocyte are presumed to be the specific cell source, as first demonstrated for locusts (Kappler et al., 1986; 1988), although this has not been confirmed in higher orders of insects (e.g., Diptera). Ecdysteroids are sequestered by developing oocytes in all insects and are present in hemolymph to a lesser or greater degree depending on whether they are required for vitellogenesis. In general, ovarian ecdysteroidogenesis proceeds in dictyopteran females in parallel to vitellogenesis (Pascual et al., 1992; Roman˜ a´ et al., 1995), but in female mosquitos and higher flies, this process is initiated by neurohormones (see Section 3.9.3.8.3) released in response to the ingestion of a blood or protein meal, respectively (Hagedorn et al., 1975; Trabalon et al., 1990). For insects, the detailed and complete biosynthetic pathway for the conversion of cholesterol to ecdysteroids is not known, but it is believed to mimic vertebrate steroidogenesis in that precursor steroid molecules shuttle between the endoplasmic reticulum and the inner mitochondrial membrane during processing (Gilbert et al., 2002; see Chapter 3.3). Key proteins and enzymes involved in the ovarian ecdysteroid biosynthetic process have been identified by genetic analysis of D. melanogaster. Females with the ecdysoneless conditional mutation appear to lack the ability to transport an intermediate to the inner mitochondrial membrane for further processing at the restrictive temperature (Warren et al., 1996), but the protein responsible for the mutant phenotype has not yet been identified. Expression of the adrenodoxin reductase (dare) gene in ovaries of D. melanogaster females is required in the vitellogenic stage of oogenesis (Buszczak et al., 1999; Freeman et al., 1999). This mitochondrial enzyme mediates the transport of electrons from NADPH to adrenodoxin, which in turn donates them to the mitochondrial cytochrome P450 enzymes responsible for steroidogenesis. The disembodied, shadow, phantom, and shade genes encode cytochrome P450 enzymes involved in the final

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hydroxylation steps of ecdysteroid biosynthesis (Gilbert et al., 2002; Warren et al., 2002; Petryk et al., 2003) (see Chapter 3.3 for details). These hydroxylases were localized within the adult ovary, and disembodied was expressed at the beginning of the vitellogenic stage of oocyte maturation (Chavez et al., 2000). Ecdysone is the major ecdysteroid secreted by A. aegypti ovaries and is converted to the more active form of 20E by the ecdysone 20monoxygenase in peripheral tissues, such as the fat body and ovaries (Hagedorn et al., 1975; Borovsky et al., 1986; Smith and Mitchell, 1986). The ovary may be a source of JH and peptide or protein messengers. The ovaries of A. aegypti synthesize in vitro physiologically significant amounts of JH III and other JH-like compounds from radiolabeled farnesoic acid, methionine, and acetate (Borovsky et al., 1994). Only one other study has investigated this phenomenon in a female insect, and under similar conditions, ovaries of D. melanogaster failed to produce JH (Richard et al., 2001). For female Diptera, ovaries with mature eggs may be the source of peptide or protein factors shown to inhibit vitellogenesis directly or indirectly (see Sections 3.9.3.8.7 and 3.9.3.8.8). Unfortunately, hemolymph titers for such factors have not been profiled in these female insects during oogenesis; thus, their physiological function remains conjectural. Several different neuropeptides have been identified in neurons and neurosecretory cells associated with oviducts (see Section 3.9.2.8), and these peptides may have paracrine and endocrine effects when released at specific times during reproduction. 3.9.2.7. Hormone Action in Female Accessory Glands

3.9.2.7.1. The role of juvenile hormone in the colleterial glands Among the insect accessory sex glands and other organs associated with fertilization

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and oviposition, the colleterial glands of female cockroaches have been studied preferentially with regard to the endocrine basis of development and biosynthetic activity. The asymmetrical left and right colleterial glands form the ootheca, the hard egg case deposited by female cockroaches. The right gland produces an 8-glucosidase, whereas the left gland produces calcium oxalate, protocatechuic acid-8-glucoside, other 8-glucosides, a diphenol oxidase, and a group of structural proteins called oothecins. When the secretions of both glands mix, the glucosidase reacts with the 8-glucosides, and the resulting phenols are oxidized to quinones, which cross-link the oothecins through phenolic bridges (see Koeppe et al., 1985). This gives the typical hard consistency of the ootheca. The first studies demonstrating the key role of CA for accessory gland development in the cockroach L. maderae were reported by B. Scharrer in the 1940s. Subsequent research in other cockroach species, P. americana and B. germanica, involving allatectomy and hormonal treatment, have demonstrated that JH is essential for the production of oothecins and the other molecules in the left colleterial gland (Figure 19). JH does not influence production of 8-glucosidase in the right colleterial gland (see Koeppe et al., 1985; Wyatt and Davey, 1996). 3.9.2.7.2. Oothecins – structural proteins regulated by juvenile hormone Oothecin synthesis in P. americana has been used as a model to investigate the effects of JH. Oothecins are composed of a 39 kDa protein, which is rich in valine and proline, and five smaller proteins (types A–E), which are rich in glycine and tyrosine. The sequence of a C type oothecin displays a number of similarities with chorion proteins of silk moths (Pau et al., 1987). Expression studies in P. americana females have shown that the first appearance of oothecin mRNAs coincides

Figure 19 The increase of protein content in the female left colleterial gland of the Blattella germanica is paralleled by the increase in production of juvenile hormone (JH) by the corpora allata (a). Protein accumulation is regulated by JH, as shown by experiments of allatectomy (CA) and subsequent treatment with 10 mg of JH III (CA þ JH) (b). (Based on data from Belles and Piulachs, 1983 and Dane`s and Piulachs, unpublished data.)

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with rising rates of JH biosynthesis, whereas allatectomy abolishes their expression. No correlation was observed with the ecdysteroid concentration in the hemolymph, whereas ovariectomy, which depresses ecdysteroid titer, did not affect oothecin production (Pau et al., 1987). Thus, ecdysteroids probably do not play a significant role in regulating oothecin expression. In a related study, 20E was shown to inhibit RNA synthesis in vitro in the left colleterial gland (Iris and Sin, 1988). This may be a pharmacological rather than physiological effect, given that the 20E dose (1.2 mg ml1) was more than two orders of magnitude higher than maximal concentrations of ecdysteroids in female P. americana (Weaver et al., 1984). 3.9.2.8. Oviposition p0560

Oviposition by female insects is regulated by the central nervous system (CNS) in response to male accessory gland factors passed during fertilization (see Chapter 1.5), oocyte maturity, and physical site characteristics. Neurotransmitters and neuropeptides, acting as such, would have a direct action in activating or inhibiting this process, but JH and ecdysteroids may modulate nervous system regulation. Diverse neurotransmitters and neuropeptides have been localized in cells of the terminal ganglion that have axons extending to different oviduct regions in insects, and their effect on oviduct muscle contractions in vitro has been documented. Tachykinins (Kwok et al., 1999), SchistoFLRFa (Kwok and Orchard, 2002), crustacean cardioactive peptide (Donini and Lange, 2002), proctolin and serotonin (Lange, 2002) stimulated such contractions, whereas octopamine and myosuppressin were inhibitory (HVFLRFa; Starratt et al., 2000). The spermatheca also are innervated from the terminal ganglion, and octopamine and Arg-Phe amide peptides alter muscle contractions in these organs (Clark and Lange, 2003). Although allatostatin-A has no demonstrated effects on insect oviducts, it greatly increases in concentration in oocytes and oviduct epithelium and nerves of the female cockroach, Diploptera punctata, during vitellogenesis, possibly functioning as an anti-JH factor in ovaries (Woodhead et al., 2003). In the cricket A. domesticus, ovipositor movements in a typical oviposition sequence are completely abolished in allatectomized females, whereas JH treatment restores that behavior (Strambi et al., 1997). Oviposition by coleopteran females is inhibited by the application of 20E antagonists, largely due to their induction of abnormal egg maturation (hyperecdysonism) (Taı¨bi et al., 2003).

Parturition hormone (PH) activity is present not only in the uterus of the tsetse fly Glossina morsitans but also in the oviducts of Bombyx and Schistocerca, as well as the ejaculatory duct of S. gregaria males (Zdarek et al., 2000). Activity thus appears to be present in the reproductive ducts of diverse insect taxa. To determine whether any of the common insect neuropeptides are capable of mimicking the effect of PH, 35 identified neuropeptides and analogs were evaluated for PH activity. Modest PH activity was observed for only high doses of proctolin and a pyrokinin analog, thus suggesting that PH is unlikely to be closely related to any of the identified neuropeptides tested. While proctolin was highly effective in stimulating contractions of the S. gregaria oviduct, the extract from the tsetse fly uterus elicited only a weak response in this bioassay. PH activity, however, was effectively mimicked with an injection of 8 bromo-cyclic GMP, suggesting a potential role for this cyclic nucleotide in mediating the PH response. Neck-ligated, pregnant females were responsive to PH, other neuropeptides and cyclic nucleotides, whereas in intact females, the brain presumably negates the effects of the exogenous compounds. 3.9.2.9. Peptide Hormones Involved in Female Reproduction

As described above, JH and ecdysteroids are considered to be the primary hormones affecting female reproduction by acting separately or coordinately to stimulate oogenesis and vitellogenesis, depending on the insect group or species. Much is known about specific neuropeptides that either stimulate or inhibit JH or ecdysteroid synthesis in insects, but little evidence points to the direct effects of peptide hormones on physiological processes associated with egg maturation (reviews: Gade et al., 1997; Klowden, 1997; De Loof et al., 2001; Na¨ ssel, 2002; Taghert and Veenstra, 2003). In effect, neuropeptides regulating the secretion of JH and ecdysteroids could be considered the ‘‘master’’ hormones, because their secretion is directed by the CNS as an integrated response to external and internal cues, such as day length and nutrient stores (see Chapter 3.2). Peptides acting as hormones or neurotransmitters have innumerable effects on behavior, ion and metabolite homeostasis, and locomotion, which are key to successful reproduction by insects (see Chapter 3.10). Only recently have comprehensive catalogs for peptide messenger genes been made available for D. melanogaster (Taghert and Veenstra, 2003) and Anopheles gambiae (Riehle et al., 2002), thanks to their respective genome projects.

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Approximately 40 different genes encoding peptide messengers are present in these dipterans, and many of these genes contain multiple copies of variant peptides. New peptide messenger genes await discovery in these insects, and future studies should elucidate the conservation of peptide messenger genes among insect orders. In addition, receptors for these peptides and elements of diverse signal transduction pathways have been revealed by bioinformatics in concert with gene expression and mutation for D. melanogaster (Taghert and Veenstra, 2003). Most importantly, elucidation of these pathways will lead to a greater understanding of how peptide messengers modulate biochemical processes and gene expression in insect cells. 3.9.2.9.1. Allatotropins The allatotropin (AT) family of structurally related peptides is so-named because the first peptide was isolated based on its stimulation of the CA in vitro to produce JH (see Chapter 3.7). In insects where vitellogenesis is JH dependent, this peptide is probably an important initiator of reproduction. The first peptide shown to have this activity was purified from head extracts of pharate adult M. sexta (Manse-AT; GFKNVEMMTARGFa; Kataoka et al., 1989). Identical peptides and genes encoding related ones have been identified in other species of Lepidoptera (Taylor et al., 1996; Oeh et al., 2000; Truesdell et al., 2000; Park et al., 2002; Abdel-latief et al., 2003). In M. sexta, three alternatively spliced mRNAs result from AT gene expression, which has been localized by in situ hybridization and immunocytochemistry to cells in the brain, frontal ganglion ventral ganglia, and midgut of different life stages (review: Elekonich and Horodyski, 2003). In P. unipuncta, expression of the AT gene was observed first in late pupae and continued during the adult stage (Truesdell et al., 2000). Related peptides and genes encoding such peptides have been identified in L. migratoria (Paemen et al., 1991), a beetle (L. decemlineata; Spittaels et al., 1996), two species of Diptera, A. aegypti (Veenstra and Costes, 1999) and A. gambiae, and in a few other invertebrates (Elekonich and Horodyski, 2003). Immunocytochemical studies with Manse-AT antisera have revealed the presence of AT-like peptides in cockroaches and two other dipterans, including D. melanogaster (Elekonich and Horodyski, 2003), but no ortholog AT gene has been identified in the D. melanogaster genome as yet (Taghert and Veenstra, 2003). In vitro assays with adult CA are routinely used to investigate the pathway of JH biosynthesis. Manse-AT stimulates the CA to produce JH I, II,

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and III, but is limited by availability of farnesoic acid, a precursor (Elekonich and Horodyski, 2003). This AT also stimulates JH secretion in vitro by larval and adult CA of other lepidopteran species and the adult CA of the honeybee, A. mellifera (Rachinsky et al., 2000) and blowfly, Phormia regina (Tu et al., 2001). Similarly, JH secretion by the CA of older sugar fed female A. aegypti is increased in vitro by this species’ AT or farnesoic acid alone, whereas both are required to activate JH synthesis in CA from newly eclosed females (Li et al., 2003). In adult insects, ATs have demonstrated effects on other processes associated with reproduction. L. migratoria AT may play a role in oviposition, as suggested by its myotropic activity on oviducts from locust and cockroach females, Leucophaea maderae, the basis for its isolation from male accessory glands (Paemen et al., 1991). Immunocytochemistry showed that AT-immunoreactivity had a widespread and sexually dimorphic distribution in the nervous system of adult locusts and cockroaches (Na¨ssel, 2002). 3.9.2.9.2. Allatostatins Three groups of neuropeptides in insects have been isolated based on their inhibition of JH secretion by CA in vitro and are known as allatostatins (ASTs). The first peptides with this bioactivity were isolated from the cockroach, D. punctata (Woodhead et al., 1989) and are now termed the A- or cockroach-type of AST (ASTA). Typically, they are short peptides with the signature terminal sequence of F/YXFGLa, and identified AST-A cDNAs and genes encode a propeptide containing 13 or 14 peptides with slight sequence variations that are posttranscriptionally cleaved and processed (Belles et al., 1999; Meyering-Vos et al., 2001). The second group, known as the B- or cricket-type of AST (AST-B), is a family of peptides with a common sequence of W(X)6Wa, e.g., GWQDLNGGWa and AWERFHGSWa, first identified in the cricket, G. bimaculatus and shown to inhibit JH biosynthesis in vitro in CA from virgin females (Lorenz et al., 1995). Related peptides were isolated first from a locust, L. migratoria, based on their inhibition of oviduct contractions (Schoofs et al., 1991). The third group or C-type of AST (AST-C) is represented by the 15 amino acid, nonamidated peptide (PEVRFRQCYFNPISCF), first discovered in M. sexta (Kramer et al., 1991), based on its inhibition of JH synthesis by CA from adult moths. All three AST types probably exist in insects, as indicated by the annotation of peptide messenger genes for D. melanogaster (Taghert and Veenstra, 2003) and A. gambiae (Riehle et al., 2002), and other reports detail the isolation of related peptides

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or characterization of encoding cDNAs or genes from a variety of insects (reviews: Bendena et al., 1999; Stay, 2000). Numerous immunocytochemical studies have examined the distribution of AST types in a great variety of insects and even different life stages and organs (Nassel, 2002). Each AST type appears to have a unique distribution in the CNS, and the presence of an AST type in the CA is dependent on whether it affects JH secretion by the CA. Interestingly, AST-A is also present in midgut endocrine cells and hemocytes (Stay, 2000), thus suggesting an even greater endocrine repertory. In general, it appears that one AST type inhibits JH secretion by CA in one or more life stages of a particular species, and the other types do not. The dipteran ASTs are exceptional in that none affect JH secretion by dipteran larval CA, but blowfly AST-A (callatostatin) inhibits JH biosynthesis by CA from cockroaches (Duve et al., 1993). Notably, peptides from all three AST types have been shown to inhibit muscle contractions in a variety of organs from different insect groups. Receptors for ASTs have been identified in a cockroach (AST-A; Auerswald et al., 2001), a silk moth (AST-A; Secher et al., 2001) and D. melanogaster, after candidate G proteincoupled receptors (GPCR) were expressed in cell systems and shown to bind AST-A (two different GPCRs; Larsen et al., 2001), AST-B (Johnson et al., 2003), and AST-C (Kreienkamp et al., 2002). In addition, ASTs have demonstrated effects on female reproduction. An AST-A inhibited vitellogenin release in vitro by fat body of cockroach females, B. germanica (Marin et al., 1996). The inhibitory effect was counteracted by the addition of mevalonolactone, thus suggesting that the AST-A inhibited synthesis of mevalonate, and hence dolichol, and therefore impairing the glycosylation and export of vitellogenin from the fat body. In vitro ecdysteroid biosynthesis by ovaries from a cricket, G. bimaculatus, was inhibited by an AST-B (Lorenz et al., 1997), the same action as prothoracicostatic peptides in the AST-B family have on larval prothoracic glands from the silk moth, B. mori (Hua et al., 1999; Chapter 3.2). Surprisingly, there are only a few reports on the in vivo effects of ASTs in female insects. After showing that bioactive AST-As were circulating in D. punctata, three synthetic AST-As were injected every 12 h for 3 days into mated females and were found to significantly decrease oocyte length and in vitro JH synthesis by the CA in treated females (Woodhead et al., 1993). Similar injections of cricket AST-A and AST-B into female crickets, G. bimaculatus, resulted in decreased body and ovary weight, eggs/ovary, and ovary ecdysteroid biosynthesis and increased vitellogenin titer in

hemolymph. The peptides had little or no effect on JH biosynthesis; however, these trends were not statistically significant when compared to controls (Lorenz et al., 1998). 3.9.2.9.3. Ovary ecdysteroidogenic hormone and neuroparsin Ovaries in female insects produce ecdysteroids, but only in Diptera do these hormones direct the increased gene expression required for vitellogenesis. In a series of classic endocrine studies on female mosquitos, the first ‘‘egg development neurosecretory hormone’’ or gonadotropin was described for insects (Lea, 1972). Almost 30 years later, this hormone was isolated from the heads of female A. aegypti, structurally characterized as an 86 residue peptide, and renamed ‘‘ovary ecdysteroidogenic hormone’’ (OEH), based on its direct stimulation of ecdysteroid synthesis by ovaries in vitro (Brown et al., 1998). Immunocytochemistry with OEH antiserum stained clusters of medial neurosecretory cells in brains, as well as other cells in the ventral ganglia and midguts of larvae and both sexes of A. gambiae and A. aegypti (Brown and Cao, 2001). Cloning of the OEH cDNA revealed the OEH prohormone and led to the bacterial expression of a recombinant OEH that was shown to have the same bioactivity in vitro and stimulated vitellogenesis and subsequent yolk deposition; this is considered to be an indirect effect of increased ovarian ecdysteroidogenesis (Brown et al., 1998). An ortholog OEH gene has been identified in another mosquito, A. gambiae, but not in D. melanogaster (Riehle et al., 2002). In Diptera, activation of ovarian ecdysteroidogenesis by OEH or other brain factors may occur through either an insulin signaling pathway (see below) or a G protein-coupled receptor (GPCR)/ cAMP pathway (Shapiro, 1983). Factors with OEH activity have been extracted from brains of higher flies (Adams and Li, 1998), and in the blowfly, Phormia regina, ovarian steroidogenesis is stimulated in vitro by cAMP analogs and preceded by a peak in ovarian cAMP levels after a protein meal (Manie`re et al., 2000). This study also showed that crude brain extracts, as well as separate extracts of the medial neurosecretory region and the rest of the brain, stimulated ovarian ecdysteroidogenesis in vitro but, surprisingly, the medial neurosecretory extracts did not elicit an increase in ovarian cAMP levels, whereas the crude brain and nonneurosecretory region extracts did. These results offer additional support for the existence of ecdysteroidogenic brain factors in female flies that act through different signaling pathways. Mosquito OEHs are closely related to the neuroparsins first isolated from the corpora cardiaca of

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L. migratoria (Girardie et al., 1987). Experiments showed that the purified native A and B forms (83 and 78 residues, respectively) administered to adult females inhibited oocyte growth, whereas injection of neuroparsin antibody alone had the opposite effect (Girardie et al., 1987). These are considered to be anti-JH effects, since JH is required for vitellogenesis in locusts, and ecdysteroids have no known role, but the mechanism for these effects remains unknown. Neuroparsins also elevated trehalose and lipid levels in hemolymph of male locusts (Moreau et al., 1988) and act as antidiuretics on rectal pads (Girardie and Fournier, 1993). As revealed by the cloning of L. migratoria neuroparsin cDNA, both neuroparsin forms are processed from a prohormone (Langueux et al., 1992). Other neuroparsins have been isolated and identified in the locust, Schistocerca gregaria, and have a similar bioactivity (Girardie et al., 1998a). More recently, four different neuroparsin transcripts/ cDNAs have been identified in S. gregaria (Janssen et al., 2001; Claeys et al., 2003); two of the transcripts were present only in brains of all life stages, whereas the other two transcripts were in the CNS, fat body, and male accessory glands and testis. Northern blot analyses showed that levels of all four transcripts changed throughout the life stages, with a significant increase in the two widely distributed ones preceding the sexual activity of males and females. Peptides for three of the transcripts have yet to be isolated, thus their role in reproduction is unknown. Locust neuroparsins and mosquito OEHs constitute a peptide family, as substantiated by an analysis of nucleotide and protein sequence databases that revealed neuroparsin related peptides in the honeybee and diverse arthropod and mollusk species (Claeys et al., 2003). This analysis further suggested that conserved features of these peptides are shared by insulin-like growth factor binding proteins, which modulate growth factor activity, in vertebrates. 3.9.2.9.4. Ovary maturing parsin A neurohormone was isolated from female locusts, L. migratoria, based on its stimulation of vitellogenesis and oocyte growth, and was thus designated ‘‘ovary maturating parsin’’ (OMP; Girardie et al., 1991). The peptide of 65 residues occurs in two isoforms, differing only at one residue position. With immunocytochemistry, OMP was localized in brain neurosecretory cells and the corpora cardiaca of locusts in all stages (Richard et al., 1994). In a later study, sets of female L. migratoria were injected daily with different amounts of OMP over 10 days after eclosion and found to have significantly increased

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hemolymph titer of ecdysteroids and oocyte length over controls (Girardie et al., 1998b). OMP has been identified in another locust, S. gregaria, and exists as isoforms: two long forms corresponding to those of L. migratoria OMP and two shorter forms found only in mature adults (Girardie et al., 1998b). Injection of mixed OMPs in female S. gregaria over 12 days similarly affected ecdysteroid and vitellogenin levels in hemolymph and stimulated oocyte growth, relative to controls (Girardie et al., 1998b). Although locust OMPs are considered ecdysteroidogenic and gonadotrophic peptides in vivo, no direct effects have been reported for OMP on isolated locust ovaries or fat body, thus, the mode of action needs further study. OMPs are known to exist in acridian Orthoptera, but not in other insects (Richard et al., 1994), as indicated by the failure to identify related peptides in the D. melanogaster and A. gambiae genome databases. 3.9.2.9.5. Short neuropeptide F, neuropeptide F, and head peptides The sequences of a great diversity of insect neuropeptides end in Arg-Phe-NH2, and three such peptides of relevance were isolated from the Colorado potato beetle, L. decemlineata, (ARGPQLRLRFa and APSLRLRFa; Cerstiaens et al., 1999) and the locust, S. gregaria (YSQVARPRFa; Schoofs et al., 2001). In a subsequent experiment, multiple injections of the beetle peptides into virgin female locusts, L. migratoria, significantly increased oocyte growth relative to controls (Cerstiaens et al., 1999). Injection of the S. gregaria peptide stimulated ovarian growth in female S. gregaria, presumably due to the increased vitellogenin levels in the peptide treated females (Schoofs et al., 2001). It has not been determined whether these effects are specific to vitellogenesis or protein synthesis in general. The two beetle peptides, along with peptides identified in other insect and arthropod species, belong to a ‘‘short neuropeptide F’’ (sNPF) family, as clearly indicated by ortholog genes in D. melanogaster (Taghert and Veenstra, 2003) and A. gambiae (Riehle et al., 2002) that encode related short peptides ending in RLRFa or RLRWa. A functional GPCR for the D. melanogaster sNPFs has been characterized and its tissue expression characterized (Mertens et al., 2002; Feng et al., 2003), but no other bioactivities are known for insect sNPFs. The ‘‘head peptides’’ first identified in A. aegypti and later shown to inhibit host seeking behavior in nonoogenic females (Brown et al., 1994) are not members of the sNPF family, as indicated by their terminus, KTRFa, and the structure of the Aedes head peptide cDNA (Stracker et al., 2002).

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Surprisingly, analysis of the D. melanogaster and A. gambiae genome databases revealed no ortholog ‘‘head peptide’’ genes (Riehle et al., 2002). The sequence of the S. gregaria peptide (probably a fragment) is more similar to the C-terminus of longer peptides (36 residues almost invariably) in the ‘‘neuropeptide F/Y’’ family of invertebrate and vertebrate peptides (Stanek et al., 2002). A functional GPCR for the D. melanogaster NPF has been characterized (Garczynski et al., 2002), and in D. melanogaster, NPF controls foraging and social behavior by larvae (Wu et al., 2003), but no bioactivities are known for adults. 3.9.2.9.6. Insulin-like peptides and the insulin signaling pathway Insulin-like peptides (ILPs) are known for only a few insect species: L. migratoria, three species of Lepidoptera, and D. melanogaster (review: Claeys et al., 2002). In all but the locust, multiple ILP genes have been identified, with B. mori having an astonishing 32 ILP genes (also known as ‘‘bombyxins’’) and D. melanogaster and A. gambiae, each with 7 ILP genes (Krieger et al., 2004). In insects, clusters of neurosecretory cells in the medial, dorsal region of brains are immunostained by insulin and ILP antisera in different life stages and species (Cao and Brown, 2001; Rulifson et al., 2002). The gut and reproductive tract may be another source of ILPs as reported for a few insect species (Montuenga et al., 1989; Iwami et al., 1996); these organs are the primary sources of insulin and related peptides in vertebrates. The presence of ILPs in hemolymph of larvae and adult silk moths has been established, and the ILP titer is much higher in males after eclosion (Satake et al., 1999). As characterized for vertebrates and invertebrates, insulin and related peptides act primarily through a receptor tyrosine kinase and an insulin receptor substrate (IRS), phosphatidylinositol 3-kinase (PI3K)/protein kinase B/Akt signaling pathway (Oldham and Hafen, 2003). For insects, the expression and function of proteins comprising this pathway have been characterized in detail only for D. melanogaster (Garofalo, 2002), and ortholog genes of the proteins were identified in A. gambiae (Riehle et al., 2002). An ever-increasing number of studies show that this pathway is a nexus for the transcriptional and translational regulation of growth and longevity in D. melanogaster and other animals (Oldham and Hafen, 2003; Tatar et al., 2003). Perturbations in any gene encoding ILPs or proteins in this pathway result in multiple and dramatic effects on not only embryonic and postembryonic development, but also oogenesis and vitellogenesis

in female D. melanogaster. Females with an ILP gene mutation had at most a single vitellogenic oocyte in each ovariole and thus a much reduced fecundity, in comparison to wild-type females (Ikeya et al., 2002). Mutations in the gene encoding the insulin receptor (IR) result in sterile females due to reduced ovariole development and yolk deposition (Chen et al., 1996). Females with an IR mutation are longer lived but deficient in JH synthesis by the CA and ovarian ecdysteroid production (Tatar et al., 2001; Tu et al., 2002). Treatment of these mutant flies with a JH analog initiates vitellogenesis and restores normal life expectancy. Vitellogenesis is blocked in females with a homozygous mutation in the IRS gene (Drummond-Barbosa and Spradling, 2001), yet another step in this key pathway. There are only a few studies of the insulin signaling pathway in other insects. In the mosquito A. aegypti, the expression pattern and phosphorylation states of the IR and PKB/Akt in ovaries were characterized through previtellogenic arrest and a gonotrophic cycle after a blood meal (Riehle and Brown, 2002, 2003). Bovine insulin stimulates ecdysteroidogenesis by A. aegypti ovaries directly in a dose-dependent manner in vitro, whereas specific inhibitors of tyrosine kinase activity and PI3K inhibit ecdysteroid production (Riehle and Brown, 1999). In Lepidoptera, a putative IR was identified in tissues from M. sexta larvae with an IR antiserum (Smith et al., 1997), and ovarian cells from three species showed high-affinity binding to a silk moth ILP, presumably to ovarian IRs recognized by an IR antiserum (Fullbright et al., 1997). Recently, the panoply of insulin-like peptide regulation in D. melanogaster, in part, has been shown to act through forkhead transcription factors and is nutrition dependent (Junger et al., 2003). The possibility also exists that insect ILPs may activate MAP kinases and G protein-coupled receptor/cAMP pathways (see Chapter 3.2), as shown for insulin related peptides in vertebrates (Hsu et al., 2002; Oldham and Hafen, 2003), and even elict cross-talk between signaling pathways. Conceptually, this activation of multiple pathways by ILPs has the potential to exhibit as profound an effect on the expression of genes required for reproduction in insects, as do JH and ecdysteroids. 3.9.2.9.7. Ovary peptides that block vitellogenesis Unrelated peptides have been isolated from mature ovaries of dipteran species based on their ability to block egg maturation. The first such peptide (YDPAPPPPPP) was extracted from the ovaries of the mosquito, A. aegypti, and shown to inhibit yolk deposition in blood-fed females (Borovsky,

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2003). Later, the direct action of this peptide was determined to be inhibition of proteolytic enzyme biosynthesis (see Chapter 4.7) in the midgut (see Chapter 4.5), and thus is known as a ‘‘trypsin modulating oostatic factor’’ (TMOF). With a TMOF antiserum, this peptide was localized to the follicular epithelium surrounding oocytes only after 24 h PBM and was not present in other tissues (Borovsky et al., 1994). This localization is supported by the characterization of a gene encoding a vitelline membrane protein, which contains TMOF sequences (Lin et al., 1993), and this protein is secreted by follicular epithelium cells. A different peptide (NPTNLH) with this same activity was purified from ovarian extracts of the flesh fly Neobellieria bullata (Bylemans et al., 1994; Borovsky et al., 1996). Injection of the hexapeptide inhibited trypsin synthesis by the midgut of liver-fed flesh fly females, resulting in a reduction of circulating vitellogenin and oocyte growth. Immunoassays determined that N. bullata TMOF-staining was localized exclusively over yolk granules in oocytes and that a 75 kDa precursor protein was present in ovary extracts (Bylemans et al., 1996). A more recent study has shown that this peptide is probably a substrate for an angiotensin converting enzyme circulating in the hemolymph of female flies, and feeding of an inhibitor of this enzyme enhanced hemolymph vitellogenin levels (Vandingenen et al., 2001). A second peptide (SIVPLGLPVPIGPIVVGPR) with a similar effect was also purified from ovarian extracts of this fly (Bylemans et al., 1995) and named ‘‘colloostatin,’’ given its sequence similarity to collagen. When administered in vivo, the peptide inhibits yolk uptake by previtellogenic oocytes and reduces the levels of circulating vitellogenin, but it does not inhibit trypsin biosynthesis in the gut. 3.9.2.9.8. Termination of vitellogenesis A few studies have shown that peptide hormones are involved in the transition from vitellogenesis to chorionogenesis so that egg maturation and ultimately oviposition can occur. Oostatic factors originating from ovaries are known for insects, and these factors may play a role in this transition through distinctly different actions, including inhibition of ovary ecdysteroidogenesis (Kelly et al., 1986; Adams and Li, 1998), than that reported for TMOF and other such ‘‘oostatic’’ factors. In dipteran females, termination of ovary ecdysteroidogenesis indirectly may end vitellogenesis, and recent studies indicate that ovarian steroidogenesis is inhibited through signaling pathways that use Ca2þ or cGMP as intermediates. In vitro ecdysteroidogenesis by vitellogenic ovaries from the blowfly,

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P. regina, was inhibited by Ca2þ ionophores or thapsigargin, whereas inhibitors of Ca2þ-calmodulin phosphodiesterases increased ecdysteroidogenesis (Manie`re et al., 2002). Analogs of cGMP inhibited in vitro steroid biosynthesis in vitellogenic ovaries from P. regina, thus correlating with the peak levels of cGMP detected in ovaries at the termination of vitellogenesis (Manie`re et al., 2003). Notably, paracrine or autocrine factors, such as nitric oxide, were implicated in this inhibition, and not brain factors, thus pointing to an even greater degree of complexity in the regulation of insect reproduction. Th adipokinetic hormone (AKH) family has key roles in regulating lipid and carbohydrate metabolism in all life stages of diverse insects (see Chapter 3.10). These peptides also are known to inhibit protein and vitellogenin synthesis and RNA synthesis in female fat body and circulate in the hemolymph of ovipositing locust females (Moshitzky and Appelbaum, 1990; Glinka et al., 1995; Kodrik and Goldsworthy, 1995). AKH regulation occurs through a GPCR, now identified in D. melanogaster and B. mori (Staubli et al., 2002), and a signaling pathway with cAMP and Ca2þ intermediates (Ga¨ de et al., 1997). A GPCR/cAMP signaling pathway is involved in the termination of vitellogenesis in mosquitos (Dittmer et al., 2003). Characterization of the gene encoding a cAMP response-element binding protein in the mosquito, A. aegypti (AaCREB) revealed signature domains for this family of transcription factors, and this gene was constitutively expressed in female fat body, where YPs are synthesized. Elicitors of the cAMP signal transduction pathway attenuated ecdysteroid stimulated YP gene expression by fat body in vitro. In cell transfection assays, AaCREB served as a potent repressor of transcription, and analysis of electrophoretic mobility shift assays detected CREB specific band-shift complexes in nuclear extracts from vitellogenic fat bodies at 24 h and 36 h post-blood meal, when YP gene expression reaches its peak then terminates. Examination of the regulatory regions of Vg and vitellogenic carboxypeptidase revealed putative CREB response elements, which bound in vitro expressed AaCREB and offers further support for its termination of YP gene expression in the fat body of this mosquito.

3.9.3. Hormones and Male Reproduction 3.9.3.1. Spermatogenesis

Insect spermatogenesis can be divided into three main steps: (1) mitotic proliferation of spermatogonia, leading to spermatocytes; (2) meiosis of

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spermatocytes, giving the spermiocytes; and (3) spermiogenesis of spermiocytes, leading to spermatozoa. As for female reproduction, ecdysteroids, JHs, and peptides have demonstrated effects on male reproductive processes. 3.9.3.1.1. Ecdysteroids and early spermatogenesis In the early steps of spermatogenesis, involving mitoses and meioses, the stimulatory role of ecdysteroids has been demonstrated in many species of Orthoptera, Hemiptera, Lepidoptera, and Diptera (Hagedorn, 1985). In R. prolixus, for example, mitotic division of spermatogonia takes place in the last larval instar in the absence of JH and in the presence of high levels of ecdysteroids. In addition, it has been shown experimentally that 20E increases the rate of mitosis in the spermatogonial cells (Dumser, 1980). Further corroboration of the importance of ecdysteroids in early spermatogenesis has been provided by Jacob (1992) who reported that fragments of testes from the rhinoceros beetle, Oryctes rhinoceros, incubated in vitro require ecdysteroids and the testis sheath to complete the mitotic and meiotic processes. The stimulatory function of ecdysteroids in early stages of spermatogenesis gives physiological sense to the discovery of ecdysteroid production in the testes of the lepidopteran Heliothis virescens (Loeb et al., 1982). These authors observed that the testis sheath of tobacco budworm larvae, when incubated in vitro, secreted ecdysteroids into the culture medium. Testes ecdysteroid production has been demonstrated for other Lepidoptera belonging to the genera Lymantria, Ostrinia, Mamestra, Leucania, and Spodoptera (see Loeb et al., 2001 and references therein). 3.9.3.1.2. The role of juvenile hormone The role of JH in spermatogenesis is less clear. Early reports suggested that JHs antagonize the stimulatory effects of ecdysteroids, especially in early stages of spermatogenesis (see Dumser, 1980; Koeppe et al., 1985; Wyatt and Davey, 1996). However, other contributions have shown that JH seems to have a stimulatory effect on late spermatogenesis, especially in diapausing species when JH simultaneously accelerates spermiogenesis and interrupts the diapause (Koeppe et al., 1985). For example, administration of a JHA to diapausing adult leafhoppers, Draeculacephala crassicornis, did not influence mitosis of spermatogonia but promoted the formation of spermatozoa (Reissig and Kamm, 1974). Results obtained in the beetle, Oryctes rhinoceros, in addition to showing that ecdysteroids are necessary in early spermatogenesis (see above), also

demonstrated that spermiogenesis took place in vitro only if an active CA pair was present in the incubation medium (Jacob, 1992). The stimulatory role of JH on spermiogenesis may be related to polyamine synthesis, especially putrescine, spermidine, and spermine, given that JH promotes the production of these compounds, at least in fat body and neural tissue of crickets (Cayre et al., 1995). Another effect of JH related to sperm physiology has been discovered by Dean and Meola (1997) in the cat flea, Ctenocephalides felis. These authors have shown that sperm transfer into the epididymis is stimulated when the fleas are exposed to a filter paper treated with JH III or JHAs. As the concentration of the JHA or the exposure time increases, the percentage of fleas that transfer sperm also increases. 3.9.3.1.3. Peptides and proteins involved in spermatogenesis In relation to ecdysteroid biosynthesis in the testes, Wagner et al. (1997) have identified a 21 amino acid peptide (ISDFDEYEPLNDADNNEVLDF) in brain extracts of the gypsy moth, L. dispar, that has ecdysteroidogenic properties. Given that the peptide induces the synthesis of ecdysteroids specifically in the testes (induction experiments using prothoracic glands were unsuccessful), it has been called ‘‘testes ecdysiotropin.’’ Other peptide or protein factors are postulated to be directly involved in the regulation of spermatogenesis. As early as 1953, C. Williams reported the occurrence of a macromolecular factor in the hemolymph of Hyalophora cecropia that stimulated spermatogenesis in spermatocysts incubated in vitro. In 1971, Kambysellis and Williams showed that 20E allowed the entry of this factor into the testis (see Hagedorn, 1985), but since then, no such factor has been isolated. More recently, spermatid differentiation in Drosophila was shown to require a peptidyl peptidase ortholog of the mammalian angiotensin converting enzyme (ACE) (Hurst et al., 2003), and ACE mRNA is found mainly in large primary spermatocytes, whereas it is not detectable in cyst cells. 3.9.3.2. Male Accessory Gland Function

The male accessory glands of insects generally are of mesodermal origin and exhibit a wide morphological diversity, from single pairs of histologically and morphologically identical tubules to multi-paired heterogeneous tubules, with multiple forms and contents (Happ, 1992). Development of male accessory glands is regulated by ecdysteroids (review: Happ, 1992), as with many other developmental processes. This section will focus on the endocrine regulation of the biosynthetic and secretory activity of the

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glands. The secretions produced by male accessory glands have a wide diversity of functions (see Chapter 1.5), and some of these secretions, when transferred to the female through copulation, may affect the process of vitellogenesis, as described below. 3.9.3.2.1. Products of the male accessory glands The male accessory glands synthesize and secrete a complex mixture of proteins, carbohydrates, lipids, and amino acids (Chen, 1984; see Chapter 1.5) that are transferred to the female during copulation. The primary function of the accessory gland products is to facilitate sperm transfer to the female. For example, these glands produce the structural proteins needed for spermatophore formation, and these spermatophore proteins may serve as nutrient resources for the female. In addition, many secretory products of the accessory glands cause physiological and behavioral changes in the mated female. These changes may be sperm related (sperm protection, storage and activation, competition with the sperm of previous males) or may alter the behavior and reproductive physiology of the mated female. The most prominent behavioral effects are the reduction of attractiveness to males and induction of refractoriness. The effects on reproductive physiology include enhancement of oocyte growth and induction of ovulation and oviposition (Chen, 1984; Gillott, 2003). In recent years, research has been focused on Drosophilid dipterans, and more than 75 accessory gland proteins have been molecularly characterized in D. melanogaster (Swanson et al., 2001; Chapter 1.5). Of these proteins, the so-called sex peptide has been the most thoroughly studied compound (see Chapter 1.5). The sex peptide of D. melanogaster is synthesized as a 55 amino acid precursor, and the processed peptide depresses sexual receptivity and enhances oviposition. Ortholog peptides have been identified in other Drosophila species (Chen, 1996; Wolfner, 1997). In D. melanogaster, sex peptide stimulates the biosynthesis of JH in vitro by the CA of the adult female (Moshitzky et al., 1996). This activity is possibly related to the results of Soller et al. (1999), which showed that the same peptide stimulates vitellogenic oocyte progression in D. melanogaster and that a JHA mimics such an effect. Interestingly, the sex peptide of D. melanogaster stimulates JH synthesis in Helicoverpa armigera (Fan et al., 1999), which suggests that this Noctuid moth may have ortholog peptides with these allatotropic properties. Male accessory glands are an important source of JHs and related metabolites (see Section 3.9.3.2.2). These JHs can be transferred to females through copulation and enhance oocyte

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growth, as shown for certain lepidopterans in which JH regulates vitellogenesis (Shirk et al., 1980; Park et al., 1998). 3.9.3.2.2. Regulation of male accessory gland function by juvenile hormone In the 1940s, Wigglesworth carried out the first studies showing the importance of CA in male accessory gland function for Rhodnius. In the cockroach, B. germanica, JH influences the growth not only of the accessory glands (Piulachs et al., 1992) (Figure 20) but also of the conglobate gland (Vilaplana et al., 1996). Further research on other insect species, mainly orthopterans, dictyopterans, lepidopterans, and dipterans, has demonstrated that allatectomy depresses to a greater or lesser degree the accumulation of gland secretions, whereas JH treatment restores the usual secretion levels (Gillott, 1996; Wyatt and Davey, 1996). In a number of species, it has been reported that the pattern of secretion accumulation in the accessory glands is parallel to that of JH concentration in the hemolymph or to that of JH synthetic rates by the CA (Wyatt and Davey, 1996). Incubation of the male accessory glands from D. melanogaster with nanomolar concentrations of JH induced a nearly threefold stimulation of protein synthesis (Yamamoto et al., 1988). Since many accessory gland proteins in D. melanogaster have been characterized at the molecular level (see above and Chapter 1.5), monitoring expression of these genes in response to JH has become easier. For example, Cho et al. (2000) have shown that the protein Mst57Dc is highly expressed after eclosion, when the titer of JH III peaks, and that in the JH deficient mutant ap56f, the levels of Mst57Dc mRNA are about 60% of those of the wild-type. Subsequent studies have monitored the effect of JH on the pattern of proteins, or even on particular proteins, present in accessory glands for species of Hemiptera, Orthoptera, and Dictyoptera (Wyatt and Davey, 1996). For example, allatectomy differentially affects the accumulation of various proteins present in the glands, whereas treatment with JH affects the protein pattern in the grasshopper, Melanoplus sanguinipes (Gillott, 1996), and in B. germanica (Belles and Piulachs, 1992). In M. sanguinipes, treatment with JH III induces the accumulation of most proteins but depresses the accumulation of two (Gillott, 1996). A later study of M. sanguinipes showed that JH acts directly on the accessory glands to promote synthesis of a specific protein, LHPI, which constitutes more than 50% of the protein content (Gillott and Gaines, 1992). Juvenile hormone could not stimulate LHPI synthesis in glands from allatectomized males, unless there was an

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Figure 20 Protein accumulation in the male accessory glands of Blattella germanica (a) is regulated by juvenile hormone (JH), as shown by allatectomy experiments (CA) and subsequent treatment with 10 mg of JH III (CA þ JH) (b). In addition, the patterns of increase in protein contents in the first days of adult life, and that of JH production by the corpora allata incubated in vitro are approximately parallel (c). (Based on data from Piulachs, M.D., Maestro, J.L., Belles, X., 1992. Juvenile hormone production and accessory reproductive gland development during sexual maturation of male Blattella germanica (L.) Dictyoptera, Blattellidae). Comp. Biochem. Physiol. 102A, 477–480; and Belles and Piulachs, unpublished data.

overnight exposure to the hormone. Cycloheximide temporally abolishes the stimulatory effect of JH on LHPI synthesis in the glands of male M. sanguinipes (Ismail et al., 1995), thus suggesting that the action of JH is mediated by protein factors involved in the transcription of LHPI. A similar priming effect of JH on protein synthesis in the accessory glands of male L. migratoria has been reported (Braun and Wyatt, 1995). It is worth noting that the protein Met of D. melanogaster, which may be involved in the action of JH action, is localized in the accessory glands and ejaculatory duct cells of adult males (Pursley et al., 2000). 3.9.3.2.3. Production of juvenile hormone by the accessory glands As mentioned above, JH is a product of the accessory gland. In fact, the first active extracts of JH were obtained by Williams in the 1950s from the abdomen of the adult male of the moth, Hyalophora cecropia, and these glands are still known to be the most copious, natural source. It was later shown that the CA of the cecropia moth produce the acids of JH I and II and that these are converted into the corresponding JHs by the male accessory glands (Shirk et al., 1983). Production of JH in the accessory glands has been reported in males of Heliothis as well (Park et al., 1998). JH production by male accessory glands may enable transfer of JH to females during copulation

to directly enhance vitellogenesis and oocyte growth or indirectly to stimulate the female’s CA. As shown for H. virescens, mating has an allatotropic effect, and JH measurements in males and females before and after copulation suggest the transfer of JH to females by the male (Park et al., 1998). Also, production or sequestration of JH by male accessory glands may be another mechanism to regulate JH hemolymph titer or biosynthesis by CA. 3.9.3.2.4. Other hormones regulating male accessory glands Hormones other than JH also may regulate the biosynthetic activity of male accessory glands. In R. prolixus males, removal of certain brain neurosecretory cells impairs protein accumulation in the accessory glands, whereas treatment with JH I only partially restores protein accumulation; a peptide extracted from R. prolixus brains stimulates protein synthesis by isolated accessory glands (Wyatt and Davey, 1996). As these results indicate, a neuropeptide and JH are required for accessory gland function. Cerebral neurosecretion and JH are involved in male accessory gland secretory activity in another blood-sucking bug, Panstrongylus megistus (Regis et al., 1987). In M. sanguinipes, cardioallatectomy inhibits the accumulation of protein LHPI to a greater degree than that provoked by allatectomy alone (Cheeseman and Gillott, 1988), which suggests again the

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participation of a neurosecretory factor in the regulation of protein secretion by the male accessory glands. Finally, 20E appears to stimulate total protein synthesis in the male accessory glands of the male lepidopterans, S. littoralis and Chilo partellus (Gillott, 1996). More detailed studies of M. sanguinipes have shown that 20E promotes protein accumulation in the accessory glands incubated in vitro (Ismail and Gillott, 1995) and that these glands incubated in vitro are able to produce ecdysteroids (Gillott and Ismail, 1995).

3.9.4. Future Directions Our understanding of how JH, ecdysteroids, and peptide hormones control various aspects of insect reproduction has advanced but also broadened greatly over the last two decades. Of particular importance is the elucidation of genetic expression heirarchies regulating ecdysteroid action at the cellular level and hormonal networks controlling reproduction at the organismal level. Despite the fact that the vitellogenic action of JH is one of the hallmarks of reproduction in most insects, its precise mechanism of action remains poorly understood. Elucidating this mechanism represents the most challenging task for future research. Genetic ecdysteroid networks governing insect development and metamorphosis have been studied in detail. Our understanding of similar networks occurring in reproduction is still limited. Future studies should take advantage of techniques for gene knockout and functional genomics to gain further insight into the regulation of female and male reproduction. Likewise, these techniques offer many advantages for research on peptide hormones involved in insect reproduction.

Acknowledgments We are grateful to Professor Mary Bownes for her kind permission to use unpublished results. We thank Dr Guoqiang Sun for his help with the manuscript, Mr. Ray E. Hardesty for editing the manuscript, Mr. Geoffrey Attardo for his help in producing figures, and Ms. Sue Gerdes for secretarial assistance.

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3.10

Hormones Controlling Homeostasis in Insects

D A Schooley, University of Nevada, Reno, NV, USA F M Horodyski, Ohio University, Athens, OH, USA G M Coast, Birkbeck College, London, UK ß 2005, Elsevier BV. All Rights Reserved.

3.10.1. Introduction 3.10.2. Hormonal Control of Energy Stores 3.10.2.1. Adipokinetic Hormone and Hypertrehalosemic Hormone 3.10.2.2. A Counter-Regulatory Hormone for AKH? 3.10.3. Hormonal Control of Water and Electrolyte Homeostasis 3.10.3.1. Molecular Basis for Urine Excretion 3.10.3.2. A Multiplicity of Peptides Regulate Diuresis and Antidiuresis in Insects 3.10.3.3. CRF-Like DH 3.10.3.4. Diuretic and Myotropic Peptides: the Kinins 3.10.3.5. Calcitonin-Like DH 3.10.3.6. CAP2b/Periviscerokinins 3.10.3.7. Arginine Vasopressin-Like Insect Diuretic Hormone 3.10.3.8. Other Peptides with Diuretic Activity 3.10.3.9. Serotonin and Other Biogenic Amines 3.10.3.10. Antidiuretic Factors Inhibiting Malpighian Tubule Secretion 3.10.3.11. Antidiuretic Factors that Promote Fluid Reabsorption in the Hindgut 3.10.3.12. Cellular Mechanisms of Action 3.10.3.13. Synergism 3.10.3.14. Possible Utility of Research on Hormonal Control of Fluid Homeostasis

3.10.1. Introduction Insects, like other animals, face a variety of challenges to their survival in the varying conditions of their environment and food availability. Many changes in their physiological status must be controlled to achieve homeostasis. In this chapter the hormonal controllers of the supply of endogenous metabolic energy, as well as the hormonal control of fluid and electrolyte balance are discussed. A review on the same subject has appeared recently, but is significantly less detailed (Ga¨de, 2004). Due to the wide variety of environmental niches inhabited by insects, the mechanisms for control of homeostasis may show considerable variation between genera, although basic underlying themes are conserved. Insects are an extremely ancient grouping of organisms, with short lifespans and frequently multiple generations per year. On completion of sequencing of the Anopheles gambiae genome, the genome and proteome of this dipteran insect were compared with that of the related dipteran Drosophila melanogaster (Zdobnov et al., 2002). One of the salient conclusions is that these two ‘‘closely related’’ dipterans show greater genetic divergence than do the genomes of the human and

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the pufferfish, although the estimated time of divergence between these two dipterans (
250 million years ago) is considerably more recent than that between the higher organisms (Zdobnov et al., 2002). This rapid rate of evolution is reflected in differences in the nature of main substrates used as metabolic fuels for flight, in considerable interspecific diversity in structure between hormones in certain families that regulate homeostasis, especially diuretic hormones, and also in differences in which family of diuretic hormones appears to dominate as the major controllers of fluid excretion between different genera. It has also become apparent that signal transduction pathways for a given family of homeostatic hormones may differ between genera. It is interesting to compare and contrast the depth of our current knowledge in this area to that when the first edition of this series was published. At that time, the sequences of only two insect neuropeptides were known: proctolin and adipokinetic hormone (AKH) from Locusta migratoria (herein called Locmi-AKH-I). Technical advances in peptide isolation and analysis made in the early 1980s were responsible for an explosive growth in the isolation and identification of new insect neuropeptides. The

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availability of synthetic peptide hormones then allowed detailed studies of their mechanism of physiological action. It has also become clear, as will be seen in this chapter, that insect peptide hormones, like their vertebrate homologs, frequently have more than one function. Well over 100 insect neuropeptides have been identified, belonging to a number of families. The majority of these were isolated on the basis of their myotropic activity, because of the ease and rapidity of bioassays for such effects. Owing to a huge literature, only those myotropic peptides are covered (certain forms of AKH and the myokinins) that are known to be important homeostatic regulators.

3.10.2. Hormonal Control of Energy Stores It is difficult to discuss this section without considering comparative endocrinology. In vertebrates the storage of fats in adipose tissue and storage of glycogen in muscles and liver are increased by high levels of insulin. Mobilization of fats from adipocytes and glycogen from liver and muscles is triggered by high levels of the counter-regulatory hormone glucagon. Glucagon acts through the cyclic AMP signal transduction pathway to activate hormone sensitive lipase in adipocytes, to activate glycogen phosphorylase and inactivate glycogen synthase in muscle and liver, and to activate gluconeogenesis and inactivate glycolysis in liver and kidney. Insulin, by activating cAMP phosphodiesterase and phosphoprotein phosphatase I, has the opposite effects on these pathways. In insects the adipokinetic-hypertrehalosemic hormones have the same role as glucagon in animals. However, a great deal of research has failed to reveal a hormone with a counter-regulatory action, although a very recent discovery by Clynen et al. (2003a) may shed light on this area. 3.10.2.1. Adipokinetic Hormone and Hypertrehalosemic Hormone

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3.10.2.1.1. Biological role of AKH Peptides in the AKH family stimulate metabolism by mobilizing energy stores in the fat body. The actions mediated by the AKH peptides are critical to supply energy to tissues, such as the flight muscles, with substrates necessary to maintain long distance flight. Depending on the insect species, either lipids, carbohydrates, proline, or a combination thereof, are released from the fat body during times of metabolic need. For example, the energy substrate released into the hemolymph of cockroaches is trehalose, a disaccharide of glucose and the major hemolymph

carbohydrate of insects (Friedman, 1985). When energy demands are high, locusts release predominantly diacylglycerols from the fat body, which are transported by lipophorins in the hemolymph (Chino and Gilbert, 1964, 1965; Soulages and Wells, 1994) (see Chapter 4.6), but also release carbohydrate under some conditions (Goldsworthy, 1969). Proline is released into the hemolymph to provide fuel for the contraction of flight muscles in the tsetse fly (Glossina morsitans; Bursell, 1963) and in coleopteran insects (De Kort et al., 1973; Ga¨ de and Auerswald, 2002). Peptides that primarily mobilize lipids in the species in which they were isolated are referred to as AKH (Mayer and Candy, 1969a), while peptides whose predominant role is to mobilize carbohydrates are known as hypertrehalosemic hormones (HrTH), since the increase in total carbohydrate is accounted for by an increase in trehalose. The adipokinetic and hypertrehalosemic functions of AKH peptides are similar to the metabolic responses induced by the vertebrate hormone, glucagon. AKH peptides are synthesized by, and released from, the cells of the glandular lobe of the corpus cardiacum (CC) (Goldsworthy et al., 1972a), an endocrine organ attached to the brain. The effect of the CC on metabolism was first described in Periplaneta americana, where injection of a CC extract into adult cockroaches caused an increased level of trehalose in the hemolymph and a decrease in fat body glycogen content (Steele, 1961). In the locusts Schistocerca gregaria and L. migratoria, injection of a CC extract into the hemolymph increases levels of diacylglycerol (Beenakkers, 1969; Mayer and Candy, 1969a), resembling the changes in lipid composition that occur during flight (Beenakkers, 1965; Mayer and Candy, 1967). This elevation of diacylglycerol reflects the oxidation of lipids as the primary fuel during prolonged flight in locusts (Weis-Fogh, 1952). The onset of flight is rapidly followed by the release of AKH into the hemolymph to mobilize energy substrates from the fat body (Cheeseman and Goldsworthy, 1979; Orchard and Lange, 1983a; Candy, 2002). The AKH concentration in the hemolymph during flight (Cheeseman and Goldsworthy, 1979; Candy, 2002) is similar to the concentration necessary to induce lipid release from the fat body in vivo (Goldsworthy et al., 1986a). In addition to the adipokinetic effect in L. migratoria, injection of a CC extract also causes an increase in hemolymph carbohydrate levels in young male locusts whose fat body contains sufficient glycogen stores (Goldsworthy, 1969). In the moth Manduca sexta, AKH stimulates glycogen breakdown in larval insects and lipid mobilization

Hormones Controlling Homeostasis in Insects

in adults (Siegert and Ziegler, 1983; Ziegler and Schulz, 1986; Ziegler et al., 1990). The similarity of AKH and HrTH was demonstrated before their structures were known; a P. americana CC extract induced an adipokinetic response in L. migratoria and a L. migratoria CC extract induced a hyperglycemic response in P. americana (Goldsworthy et al., 1972a). The differences in the metabolic responses reflect the strategy of the insect in the nature of the fuel stored in the fat body. An adipokinetic hormone was first identified in L. migratoria (Stone et al., 1976). The structural similarity of the L. migratoria and P. americana peptides was confirmed when three groups isolated two octapeptides in the same year from P. americana based on myotropic assays (Baumann and Penzlin, 1984; Scarborough et al., 1984; Witten et al., 1984). However, Scarborough et al. (1984) also showed that the less abundant peptide from P. americana, Peram-CAH-II (Figure 1) has a very significant sequence identity to glucagon (4 of 8 residues are identical; 2 others are highly conserved), and that both peptides would elevate hemolymph carbohydrate in an assay where trehalose is hydrolyzed to glucose. Later Siegert and Mordue (1986a) isolated these factors as specific hypertrehalosemic agents from cockroach. Because of the functional relatedness of AKH peptides, heterologous bioassays, such as the lipid mobilization assay in L. migratoria and carbohydrate mobilization assay in P. americana, are often used for the isolation of metabolic neuropeptides in a large number of insect species (Ga¨ de, 1980). The biochemical effect of a CC extract in the P. americana fat body is the activation of glycogen phosphorylase (Steele, 1963). Similarly in L. migratoria, glycogen phosphorylase is activated by a CC extract (Van Marrewijk et al., 1980) or by synthetic AKH (Ga¨ de, 1981). The effect of AKH on lipid mobilization in M. sexta is mediated by the activation of triacylglycerol lipase (TAG lipase) which converts TAG into diacylglycerol, which is subsequently released into the hemolymph (Arrese et al., 1996). Injection of Locmi-AKH I into S. gregaria resulted in a twofold increase of TAG lipase in the fat body (Ogoyi et al., 1998). As is common among neuropeptides, AKH possesses numerous biological activities in addition to its well-known metabolic functions. Some of these actions include the acceleration of heart rate in P. americana (Baumann and Gersch, 1982; Scarborough et al., 1984) and Blaberus discoidalis (Keeley et al., 1991), the stimulation of myotropic contractions in P. americana (O’Shea et al., 1984; Witten et al., 1984), the inhibition of protein synthesis

495

Figure 1 Alignment of sequences of the known AKH/HrTH from the Insecta, from a sample of 39 peptides with a unique primary structure. Peptide sequences were compiled largely from Ga¨de et al. (1997) with the inclusion of additional unique sequences found since 1997 (Ga¨de and Kellner, 1999; Siegert, 1999; Kodrı´ k et al., 2000; Ko¨llisch et al., 2000; Siegert et al., 2000; Lorenz et al., 2001; Ga¨de et al., 2003). The sequence of one peptide from Carausius morosus (Carmo-HrTHI*) contains a hexose modification on the Trp8 residue (Ga¨de et al., 1992), and an additional peptide from Platypleura capensis (Placa-HrTHIy) contains a modification that has not been characterized (Ga¨de and Janssens, 1994). Species names are as given in the Swiss-Prot database as recommended previously (Coast et al., 2002). Those not defined in the text are: Anaim, Anax imperator; Bladi, Blaberus discoidalis; Declu, Decapotoma lunata; Emppe, Empusa pennata; Erysi, Erythemis simplicicollis; Grybi, Gryllus bimaculatus; Libau, Libellula auripennis; Micvi, Microhodotermes viator; Onyay, Onymacris aygulus; Phote, Phormia terranova; Phymo, Phymateus morbillosus; Phyle, Phymateus leprosus; Polae, Polyphaga aegyptiaca; Psein, Pseudagrion inconspicuum; Pyrap, Pyrrhocoris apterus; Rommi, Romalea microptera; Scade, Scarabaeus deludens; Tenar, Tenthredo arcuata; Vanca, Vanessa cardui. The 15 N-terminal residues of glucagon are shown (16–29 deleted); Gln3 is conserved in all AKH and the motif Thr-Phe-Thr from glucagon is 100% conserved in nine sequences, and Ser8 and Asp9 of glucagon are conserved in certain AKH sequences and are conservative substitutions in others. Tyr10 is a conservative substitution for the completely conserved Trp.

496 Hormones Controlling Homeostasis in Insects

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(Carlisle and Loughton, 1979; Cusinato et al., 1991) and RNA synthesis (Kodrik and Goldsworthy, 1995), the inhibition of lipid synthesis (Gokuldas et al., 1988), the stimulation of heme synthesis in B. discoidalis (Keeley et al., 1991), a neuromodulatory effect in P. americana (Wicher et al., 1994), the stimulation of locomotory activity in M. sexta (Milde et al., 1995) and Pyrrhoccoris apterus (Socha et al., 1999; Kodrı´k et al., 2000), and a lipogenic effect in which lipids are translocated from the hemolymph to the fat body in P. americana (Oguri and Steele, 2003). Although AKH is widely viewed as an energy mobilization hormone synthesized by and released from the CC, effects on the nervous system have been documented. This was first shown by demonstrating that neurohormone D, an HrTH neuropeptide in P. americana, elicits an increase in the spontaneous spike frequency in DUM neurons of the cockroach terminal ganglion (Wicher et al., 1994). This action of neurohormone D on Ca2þ currents is through the upregulation of the mid/low voltage activated (M-LVA) Ca2þ current by the actions of cAMP and protein kinase A (PKA), suggesting a role for phosphorylation of the channel protein by PKA (Wicher, 2001). The contribution of protein kinase C in the regulation of Ca2þ influx through voltage activated Ca2þ channels was also documented. Injection of Manse-AKH into the mesothoracic neuropil increases motor activity in M. sexta, similar to the effects of octopamine (Milde et al., 1995). A stimulation of locomotory activity in the firebug, P. apterus was observed after injection of Locmi-AKH I or the endogenous peptide, PyrapAKH (Socha et al., 1999; Kodrı´k et al., 2000). The mechanism for the peptide’s action on the nervous system was not determined, but dispersal activity is associated with starvation and the need to mobilize energy reserves from the fat body (Socha et al., 1998). Thus, in P. apterus, the neuromodulatory function of AKH integrates behavior with the metabolic needs of the insect (Socha et al., 1999). Some investigators reported the presence of Locmi-AKH I in the brain (Moshitzky et al., 1978; Bray et al., 1993). Using mass spectrometric (MS) analysis, none of the Locmi-AKHs were detected in the pars intercerebralis (PI), but a material with the Mr of Locmi-HrTH (Siegert, 1999) was present in the PI and in the CC (Clynen et al., 2001). The interaction of AKH with both the humoral and cellular immune system was demonstrated in L. migratoria (Goldsworthy et al., 2002, 2003a). One component of the insect defense response to wounding and infection is the activation of phenoloxidase via a serine proteinase cascade triggered

by the recognition of bacterial and fungal cell wall components resulting in the production of toxic quinones (Gillespie et al., 1997). Injection of Locmi-AKH I or II enhanced the activation of phenoloxidase in response to a challenge with laminarin (a b-1,3-glucan) and activated phenoloxidase in the presence of bacterial lipopolysaccharide (LPS) (Goldsworthy et al., 2002). The different potencies of Locmi-AKH I and II in phenoloxidase activation are consistent with their relative potencies in the locust lipid mobilization assay (Goldsworthy et al., 1986b), suggesting a direct link between the immune and lipid mobilization responses (Goldsworthy et al., 2002). The enhancement of phenoloxidase activation by AKH is age dependent, and restricted to the mature adult stage (Mullen and Goldsworthy, 2003). This time dependence correlates with the lipid mobilization response to AKH (Mwangi and Goldsworthy, 1977a) and to the concentration of apolipophorin III (ApoLp-III ) in the hemolymph (Mwangi and Goldsworthy, 1977b; Mullen and Goldsworthy, 2003). In addition to the role of ApoLp-III in lipid metabolism, the lipid associated form of ApoLp-III stimulates antimicrobial activity in the hemolymph of Galleria mellonella (Wiesner et al., 1997; Detloff et al., 2001). These data suggest that AKH may be exerting its effect on phenoloxidase activation in part through its effects on ApoLpIII metabolism (Mullen and Goldsworthy, 2003). An effect on phenoloxidase activation by LPS in the absence of AKH was observed in starved locusts, presumably because of the effects of starvation on lipid mobilization (Goldsworthy et al., 2003a). An important component of the cellular immune response is the formation of nodules by hemocyte aggregation to engulf or entrap foreign bodies (Lavine and Strand, 2002). AKH increases nodule formation when coinjected with LPS (Goldsworthy et al., 2003a). Unlike the effects of AKH on phenoloxidase activation, which is specific to mature locusts (Goldsworthy et al., 2002), its effects on nodule formation are apparent from the fifth instar nymphal stage through the mature adult stage. AKH may be exerting its effect on the humoral immune system (activation of prophenoloxidase) and the cellular immune system (nodule formation) via separate mechanisms, since they exhibit differing sensitivities to pharmacological agents and show different correlations to the nutritional state and lipid composition (Goldsworthy et al., 2003b). 3.10.2.1.2. Structural features of AKH Evidence for multiple AKHs within a species was first found in locusts (Carlsen et al., 1979), and has been extended to include many insect species (Ga¨de

Hormones Controlling Homeostasis in Insects

et al., 1997), with three biologically active peptides present in some insects (Oudejans et al., 1991; Siegert et al., 2000; Ko¨ llisch et al., 2003). Each AKH is derived from a unique mRNA (Schulz-Aellen et al., 1989; Noyes and Schaffer, 1990; Fischer-Lougheed et al., 1993; Bogerd et al., 1995). In L. migratoria, a fourth member of the AKH family (Locmi-HrTH) was characterized based on its hypertrehalosemic activity in P. americana, but no function has yet been identified in L. migratoria (Siegert, 1999). The three Locmi-AKHs differ in amount, with LocmiAKH I being the most abundant and Locmi-AKH III the least abundant (Oudejans et al., 1993; Clynen et al., 2001). One rationale for multiplicity of peptides is their varying potencies in different biological assays, which may enable the fine tuning of an energy-demanding response such as long-distance flight (Vroemen et al., 1998a). Locmi-AKH I and II differ in potency. Locmi-AKH I is more effective than Locmi-AKH II in the lipid mobilization assay, the ability to alter circulating lipoproteins, and the ability to activate glycogen phosphorylase, while Locmi-AKH II is more effective in increasing cAMP levels in the fat body (Orchard and Lange, 1983a; Goldsworthy et al., 1986b). These data suggest that Locmi-AKH I is the predominant hyperlipemic hormone, and Locmi-AKH II is largely responsible for carbohydrate mobilization. Locusts employ carbohydrate as an energy source for the initial stages of flight and lipid for more prolonged flight (Weis-Fogh, 1952; Mayer and Candy, 1969b; Jutsum and Goldsworthy, 1976), suggesting that Locmi-AKH II is most important at the onset of flight and Locmi-AKH I assumes a major role in prolonged flight activity (Vroemen et al., 1997). Locmi-AKH III is present at low levels and seems to have a high turnover, suggesting it may be a modulatory entity that provides the locust with energy when not flying. The peptides from the AKH family are among the most thoroughly characterized insect peptides with respect to their structural diversity (Ga¨de et al., 1997). At least 38 distinct peptides have been structurally characterized from at least 90 species that include representatives from most insect orders (Figure 1). A distinct sequence of an additional family member, the red pigment-concentrating hormone (Panbo-RPCH), long thought to be unique to crustaceans (Fernlund and Josefsson, 1972), has recently been found in insects (Ga¨de et al., 2003). RPCH has been shown to induce hyperlipemia in locusts (Herman et al., 1977; Mordue and Stone, 1977). In contrast to the insect peptides whose roles include the mobilization of metabolic stores, the role of RPCH in crustaceans is to induce aggregation of pigment

497

granules in chromatophores (Rao, 2001). Thus, these orthologous peptides found in insects and crustacea do not retain a common function. Although AKH-like immunoreactivity exists in invertebrate phyla other than the Arthropoda (Schooneveld et al., 1987), confirmation of the presence of related peptides is lacking. The most compelling evidence thus far for an AKH related peptide in a nonarthropod is the detection of Locmi-AKH I-like immunoreactivity in the nematode Panagrellus redivivus and the induction by a peptide fraction of an adipokinetic response in S. gregaria and a hypertrehalosemic response in P. americana (Davenport et al., 1991). All AKH precursor proteins share a common organization; an N-terminal signal peptide followed by AKH and the AKH precursor-related peptide (APRP) (Martı´nez-Pe´ rez et al., 2002). The organization of the RPCH precursor protein (Linck et al., 1993) is similar to that of AKH/HrTH peptides, supporting the hypothesis that they are derived from a common ancestor (Martı´nez-Pe´ rez et al., 2002). Close examination of the precursor structures and gene organization of AKH/RPCH peptides with that of the APGWamide peptide family of mollusks revealed similarities including a common overall architecture of the precursor protein, a common predicted secondary structure, and similar position of introns, suggesting that they could have arisen from a common ancestral gene or from recombination of an ancestral AKH/RPCH-like gene with an APGWamide-like gene (Martı´nez-Pe´ rez et al., 2002). While significant structural variability was observed when comparing AKH peptides sequenced in different insect species, several common characteristics are defining features of this family (Ga¨ de et al., 1997) (Figure 1). The sizes of AKHs range from 8 to 11 amino acids. All AKHs are blocked at their N-terminus with a pyroglutamate residue, and all are amidated at their C-terminus with one exception (Ko¨ llisch et al., 2000). An aromatic amino acid is present in each AKH at position 4, and Trp is conserved at position 8. The most critical residues for bioactivity are probably the most prevalent (pGlu1, Phe4, Trp8) and might be involved in binding to the receptor, stability of the peptide, or formation of an intrapeptide interaction stabilizing the conformation needed to achieve optimal activity (Hayes and Keeley, 1990; Ga¨ de, 1992). For example, the conserved Trp8 was proposed to form a hydrogen bond with Ser5 or Phe4 (Hayes and Keeley, 1990), and the hydrophobic nature and relative spacing of residues 2, 4, and 8 may be important in determining receptor binding.

498 Hormones Controlling Homeostasis in Insects

The biological activity of neuropeptides is mediated by their interaction with a membrane bound receptor on the target cell; most then alter the concentration of second messengers that result in biochemical changes and ultimately lead to a physiological response. Small peptides, such as AKH, are flexible structures that adopt many conformations in solution, and their potency is related to their structural conformation upon interaction with the receptor (Nachman et al., 1993). The identification of AKH receptors and their expression allows the direct assay of peptide (or analog) binding to the receptor and subsequent response of the target cell to identify the critical structural features of the peptides that are required for receptor binding and activation. However, considerable information on the importance of individual amino acids or parts of the peptide for binding to the receptor has been acquired using bioassays in structure–activity studies in several insect species, but these results are also influenced by such factors as the relative stability of the peptides in vivo. The overall conclusion that can be drawn from structure–activity studies is that the endogenous peptides are usually the most potent in the respective bioassays, and coevolution of the peptide and the receptor took place to achieve optimal binding (Hayes and Keeley, 1990; Fox and Reynolds, 1991a). Stone et al. (1978) predicted that many members of the AKH/RPCH family have the potential to adopt a b-turn conformation. While AKH peptides do not form an ordered structure in aqueous solution, the presence of sodium dodecylsulfate (SDS) causes a change in the circular dichroism (CD) spectra that is characteristic of a b-turn (Goldsworthy and Wheeler, 1989; Cusinato et al., 1998). However, the CD spectra of Locmi-AKH II and ManseAKH, peptides that lack a proline residue in position 6, are not affected by the presence of SDS and lack the capacity to form a b-turn. The capacity of some AKH peptides to form a b-turn between residues 4 and 8 was also confirmed using nuclear magnetic resonance (NMR) spectroscopy (Zubrzycki and Ga¨ de, 1994; Zubrzycki and Ga¨de, 1999; Nair et al., 2001). It was suggested that the activity of AKH in the L. migratoria lipid mobilization assay is correlated with the ability to form a b-structure in SDS micelles, which in turn quantitatively affects its interactions with the receptor (Goldsworthy et al., 1997; Cusinato et al., 1998). Locmi-AKH II and ManseAKH, which lack the ability to form a b-structure, exhibit low potency in the L. migratoria lipid mobilization assay (Cusinato et al., 1998). Analogs with substitutions in residues 6–8 of AKHs exhibit reduced potency since the substitutions apparently

hinder the formation of a b-turn (Lee et al., 1996). However, the capacity to form a b-turn does not correlate with potency when assayed for inhibition of acetate uptake into fat body in vitro, since Acheta domesticus AKH and Locmi-AKH II are equally active using this assay (Cusinato et al., 1998). This pleiotropic response may reflect the heterogeneity of receptors, which exhibit differing structural preferences for their ligand. Structural analysis of AKH peptides from Melolontha melolontha (Melme-CC), Tenebrio molitor (Tenmo-HrTH), and Decapotoma lunata (Declu-CC) using NMR supports the hypothesis that the b-turn is important for receptor binding in the L. migratoria lipid mobilization assay (Nair et al., 2001). The Asn residue at position 7 (Asn7), and the Phe residue at position 4 (Phe4) are essential for a potent response, and it was shown that the Asn7 projects outward from the b-turn, and the orientation of Asn7 and the tightness of the b-turn are influenced by the aromatic residue at position 4. Data on the biological activity of a large number of natural AKHs and synthetic analogs in the L. migratoria lipid mobilization assay and acetate uptake assay were compiled to attempt to construct quantitative structure–activity relationships (QSAR) (Lee et al., 2000). Little new understanding was gained in this study since the peptides used were not designed with QSAR in mind, but this approach might be used to understand the requirements of peptide–receptor interactions and to predict sequences that possess improved biological potency as used successfully with substance P (Norinder et al., 1997). Structure–activity studies were also carried out using the activation of glycogen phosphorylase in M. sexta fat body to assay several naturally occurring AKH peptides and synthetic analogs (Ziegler et al., 1991, 1998). Manse-AKH lacks the ability to form a b-turn (Goldsworthy and Wheeler, 1989), and among the peptides tested, the most inactive were the ones with the highest probability of forming a b-turn (Ziegler et al., 1991, 1998). The substitution of Ser6 of Manse-AKH with Pro, which favors the formation of a b-turn, drastically reduced its abilities to activate glycogen phosphorylase and to bind to the receptor in a fat body membrane preparation. This indicates that the M. sexta AKH receptor does not bind peptides that contain a b-turn (Ziegler et al., 1998). Removal of pGlu1 from Locmi-AKH I abolishes activity (Ga¨de, 1990), but replacement of pGlu1 by an unblocked amino acid partially restored lipid mobilization activity in vivo to varying degrees depending on the identity of the substituted amino acid, and activity was further enhanced by N-terminal blockage (Lee et al., 1997). Replacement of pGlu1 in

Hormones Controlling Homeostasis in Insects

Manse-AKH, however, did not restore potency in a bioassay measuring activity of glycogen phosphorylase unless the N-terminus was blocked (Herman et al., 1977; Ziegler et al., 1998). These data imply that pGlu1 is not absolutely essential for activity, and that one of the roles of the blocked N-terminus might be to impart increased stability to the peptide by preventing aminopeptidase attack (Lee et al., 1997; Ziegler et al., 1998). 3.10.2.1.3. AKH synthesis and release Like most neuropeptides, AKH is derived by processing of a larger precursor protein (Hekimi and O’Shea, 1987). The steps involved in AKH synthesis have been most thoroughly characterized in S. gregaria (O’Shea and Rayne, 1992), and subsequent studies in other insects have shown that the basic features of AKH biosynthesis are conserved. AKH precursor proteins share a similar organization, an N-terminal signal peptide, followed by AKH and the AKH precursor related peptide (APRP) (O’Shea and Rayne, 1992), as first shown in S. gregaria (Schulz-Aellen et al., 1989) and M. sexta (Bradfield and Keeley, 1989). The locust AKH precursor exists as a dimer formed by oxidation of the Cys residues present in the APRP. This occurs prior to proteolytic processing in the trans-Golgi at basic amino acid residues and C-terminal amidation (Rayne and O’Shea, 1994). The dimeric structure of the precursor might confer a conformation that facilitates or allows the correct processing of the precursor. Since multiple AKHs (Diederen et al., 1987; Hekimi et al., 1991) and their mRNAs (Bogerd et al., 1995) are present in the same glandular cells, random formation of dimeric precursors gives rise to both homodimers and heterodimers (Hekimi et al., 1991; Huybrechts et al., 2002). Further processing of L. migratoria APRPs from the AKH I and AKH II precursors takes place to yield additional peptides designated the adipokinetic hormone joining peptides (AKH-JP I and AKH-JP II), but their release was not demonstrated (Baggerman et al., 2002). Biological roles for the APRPs or the AKH-JPs have not been found (Oudejans et al., 1991; Hatle and Spring, 1999; Baggerman et al., 2002). Relative peptide levels in the glandular cells are controlled by multiple mechanisms. The 4.5:1 ratio of AKH I to AKH II present in the glandular cells (Hekimi et al., 1991) is regulated by a 1.7:1 ratio of AKH I to AKH II mRNA and the more efficient translation of AKH I mRNA (Fischer-Lougheed et al., 1993). Antisera specific to AKH I or AKH II have demonstrated their colocalization in the same secretory granules in the glandular cells of the CC (Diederen et al., 1987). Since an AKH III-specific antiserum is

499

not available, the colocalization of each APRP implies that AKH III is also colocalized with AKH I and II (Harthoorn et al., 1999). The amount of AKHs increase dramatically in the CC during development (Siegert and Mordue, 1986b), due to their continued synthesis (Fischer-Lougheed et al., 1993; Oudejans et al., 1993) and to the increase in the number of cells in the glandular lobe of the CC (Kirschenbaum and O’Shea, 1993). The continued synthesis of AKH generates a large pool of secretory granules in the glandular cells (Diederen et al., 1992), and newly synthesized AKH is preferentially released over AKHs stored in older secretory granules (Sharp-Baker et al., 1995), which constitutes a pool of peptides that cannot be released (Sharp-Baker et al., 1996; Harthoorn et al., 2002). Therefore, only a small percentage of the total AKH present in the glandular cells is released during flight (Cheeseman et al., 1976) or upon stimulation by CCAP (Harthoorn et al., 2002). The AKHs are released in the same proportion as their levels in the CC (Harthoorn et al., 1999), and the continuous synthesis of AKH is required for the secretion of peptides from the glandular cells in response to metabolic need (Harthoorn et al., 2002). In S. gregaria and L. migratoria, the hemolymph trehalose levels during flight are about 50% that of preflight values (Mayer and Candy, 1969b; Jutsum and Goldsworthy, 1976), and injection of a high concentration of trehalose prevents AKH release assayed by quantifying lipid mobilizing activity in the hemolymph (Cheeseman et al., 1976). Trehalose and glucose exert a direct action on the glandular cells of the CC, since high trehalose concentrations decreased both spontaneous AKH release and AKH release induced by the neuropeptide Locmi-TK I, 3-isobutyl-l-methylxanthine (IBMX), or high potassium concentrations in vitro (Passier et al., 1997). It was suggested that trehalose exerts this effect, in part, after its conversion to glucose. The decrease in trehalose concentration in response to the energy demands of flight relieves this inhibition, and is one of the factors that contributes to AKH release that is observed during flight (Cheeseman and Goldsworthy, 1979; Orchard and Lange, 1983b). It was also suggested that high levels of diacylglycerol resulting from mobilization of lipid energy stores may exert negative feedback on AKH release (Cheeseman and Goldsworthy, 1979). The neural and hormonal factors that modulate AKH release have been most thoroughly characterized in L. migratoria (Vullings et al., 1999). Implanted CC do not show the ultrastructural signs of enhanced AKH release during flight, suggesting that AKH secretion is under neural control by cells

p0125

500 Hormones Controlling Homeostasis in Insects

that make direct contact with the glandular lobe cells (Rademakers, 1977a). Neuroanatomical studies defined the secretomotor neurons in the lateral part of the protocerebrum that project through the nervi corporis cardiaci II (NCC II) to innervate the glandular lobe of the CC (GCC) and make synaptic contact with the AKH cells, and the axon terminals of the GCC are derived solely from the secretomotor cells (Rademakers, 1977b; Konings et al., 1989). Electrical stimulation of the NCC II resulted in the release of AKH I and II from the GCC and was accompanied by an increase in cAMP levels in cells of the GCC (Orchard and Loughton, 1981; Orchard and Lange, 1983a). While stimulation of the NCC I had no direct effect on AKH release, an enhancement of the NCC II-stimulated release of AKH from the CC was observed (Orchard and Loughton, 1981). Hormonally mediated lipid mobilization during flight is abolished in locusts in which the NCC I and NCC II were severed (Goldsworthy et al., 1972b). Together, these data indicate that the secretomotor neurons are involved in the control of AKH release, and compounds present in the NCC I modulate AKH secretion. An antiserum raised against locustatachykinin I (Locmi-TK I), a member of a family of structurally related neuropeptides (Schoofs et al., 1993), stained a subset of secretomotor neurons projecting to the GCC and made synaptoid contacts with AKHimmunoreactive glandular cells, suggesting a role for a Locmi-TK I-like peptide in AKH release (Na¨ ssel et al., 1995). Indeed, Locmi-TK I and II induced AKH release from the CC in vitro and increased cAMP levels in the GCC, but an effect on AKH release required concentrations in the 50–200 mM range (Na¨ssel et al., 1995, 1999). Furthermore, four Locmi-TK isoforms were identified in L. migratoria CC extracts (Na¨ssel et al., 1999). The anatomical features of the Locmi-TK-containing secretomotor neurons suggest that these peptides might act on synaptic receptors, and the high concentration of peptide required for AKH release is consistent with this hypothesis, particularly since synaptic receptors might be less accessible for an in vitro effect and thus would require higher peptide concentrations for an effect to be observed. SchistoFLRFamide is a member of a large family of neuropeptides known as the FMRFamide-related peptides (FaRPs) that are widely found in insects (Orchard et al., 2001). Antisera to FMRFamide and SchistoFLRFamide label a subset of secretomotor neurons that are distinct from the Locmi-TKcontaining cells which make synaptoid contacts with glandular cells of the CC (Vullings et al., 1998). FMRFamide and SchistoFLRFamide have no effect on the spontaneous release of AKH in vitro, but

reduced IBMX-induced AKH release at a 10 mM peptide concentration. Like the case with Locmi-TK neuropeptides, the high concentration of FaRPs required for an effect is consistent with a direct supply of peptides on the glandular cells. Thus, FaRPs are inhibitory neuromodulators that act to fine-tune the release of AKH to meet the energetic demands of the flying animal (Vullings et al., 1998). A direct approach was used to isolate a compound, the neuropeptide CCAP, from a S. gregaria brain extract that potently stimulates release of AKH from L. migratoria and S. gregaria CC in vitro (Veelaert et al., 1997). CCAP is a multifunctional neuropeptide and is best known for its stimulatory activity on heartbeat (Tublitz and Truman, 1985) and its effect as trigger for ecdysis behavior (Gammie and Truman, 1997; see Chapter 3.1). Unlike the Locmi-TK and SchistoFLRFamide-like peptides, there are no CCAP containing fibers in the GCC (Dircksen and Homberg, 1995; Veelaert et al., 1997); CCAP is active in the nM range indicating that it may act as a neurohormonal releasing factor for AKH (Veelaert et al., 1997). Flight activity increases the octopamine levels in the hemolymph of S. gregaria (Goosey and Candy, 1980), and in L. migratoria, octopamine was shown to stimulate AKH release from the CC in the presence of IBMX (Pannabecker and Orchard, 1986). It was later found that in this experimental design, octopamine potentiates the stimulatory effect of IBMX mediated cAMP elevation on AKH release, and has no effect on its own (Passier et al., 1995). This suggests that octopamine has a neurohormonal role in modulating AKH release (Veelaert et al., 1997). 3.10.2.1.4. AKH degradation After secretion of AKHs into the hemolymph, their degradation will have a significant effect on peptide levels, and the differential rates of degradation will affect the ratios of peptides and, ultimately, the physiological response. AKHs are blocked at both termini, and the initial step in their degradation is by an endopeptidase that yields inactive peptide fragments (Siegert and Mordue, 1987; Fox and Reynolds, 1991b; Rayne and O’Shea, 1992). The endopeptidase has been localized to the external surface of several tissues (Rayne and O’Shea, 1992), including the Malpighian tubules (Baumann and Penzlin, 1987; Siegert and Mordue, 1987), but in M. sexta, AKH is cleaved by an enzyme circulating in the hemolymph (Fox and Reynolds, 1991b). The half-life of AKH I in L. migratoria was initially determined to be 24 min by quantifying the disappearance of AKH from the hemolymph subsequent to its release using an in vivo bioassay (Cheeseman

Hormones Controlling Homeostasis in Insects

et al., 1976). Subsequently, a half-life of 30 min was determined for AKH I and II in S. gregaria by measuring the disappearance of AKH after injecting a large, nonphysiological dose (Rayne and O’Shea, 1992). The synthesis of high-specific activity tritiated AKHs allowed the quantification of the half-lives of AKH I, II, and III in L. migratoria after injecting a physiological dose of 1 pmol (Oudejans et al., 1996). This study clearly demonstrated that each AKH has a different rate of breakdown, with AKH III being significantly less stable in the hemolymph. The halflives of AKH I, II, and III at rest are 51, 40, and 5 min, respectively, and AKH I and III are degraded more rapidly during flight, suggesting that more than one protease is involved in AKH degradation. In contrast to the relatively long half-lives of Locmi-AKH I and II, injected Gryllus bimaculatus AKH has a half-life of about 3 min and is degraded by enzymes released from hemocytes (Woodring et al., 2002).

p0165

3.10.2.1.5. Signal transduction for AKH There is substantial evidence from a number of systems that inositol-1,4,5-trisphosphate (IP3) serves as the second messenger to transduce the hormonal signal of AKH/HrTH neuropeptides in the fat body and, in some cases, cAMP is also involved (Van der Horst et al., 1997; Ga¨ de and Auerswald, 2003). In L. migratoria, the involvement of cAMP in AKH mediated lipid mobilization was shown by demonstrating the accumulation of cAMP in the fat body following an injection of a CC extract, and the increased hemolymph lipid concentration after treatment with dibutyryl-cAMP (db-cAMP) (Ga¨ de and Holwerda, 1976). A similar effect of a CC extract on cAMP levels was also shown in vitro in S. gregaria (Spencer and Candy, 1976). The effect of the CC extract on cAMP levels was shown to be due to AKH (Ga¨ de, 1979; Goldsworthy et al., 1986b). Although the mobilization of lipids is a crucial role of AKH in L. migratoria, the mechanism of AKH action in the target cell has been studied most extensively using glycogen phosphorylase activation as the assay for AKH activity (Van Marrewijk et al., 1980). The potencies of each AKH in cAMP elevation and glycogen phosphorylase activation differed when assayed at physiological levels, decreasing in the order AKH III > AKH II > AKH I (Vroemen et al., 1995a). These differences in potency support the hypothesis that the action of AKH II is more directed to carbohydrate metabolism than that of AKH I (Orchard and Lange, 1983a). The increase in cAMP levels is mediated by the G protein Gs, since cholera toxin, an irreversible activator of Gs, enhanced cAMP levels and glycogen phosphorylase activity (Vroemen et al., 1995a). Pertussis toxin,

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an irreversible inhibitor of Gi, had no effect on cAMP levels or glycogen phosphorylase activation. In L. migratoria, the uptake of Ca2þ into the fat body was stimulated by the presence of AKH, and the effects of AKH on diacylglycerol levels and glycogen phosphorylase activation are dependent on extracellular Ca2þ and can be mimicked by the calcium ionophore A23187 (Van Marrewijk et al., 1991). A similar Ca2þ dependence for AKH mediated lipid mobilization was demonstrated in vitro (Lum and Chino, 1990; Wang et al., 1990). In addition, the release of Ca2þ from intracellular stores leads to the subsequent influx of Ca2þ into the cell (Van Marrewijk et al., 1993) by a process known as capacitative Ca2þ entry (Berridge, 1995), confirming a requirement for both intracellular and extracellular Ca2þ for the full effect of AKH on the fat body. Although each of the three AKHs increase Ca2þ uptake with similar potency, the three AKHs also enhance the efflux of Ca2þ into the fat body cytosol, but the increase in efflux increased in the order AKH I < AKH II < AKH III (Vroemen et al., 1995b), so the effect of AKH I on intracellular Ca2þ levels that result from flux across the cell membrane is the greatest of the three AKHs. The mobilization of intracellular Ca2þ by each of the AKHs in locusts is mediated by the stimulation of inositol phosphate (InsPn) formation, most notably IP3 (Stagg and Candy, 1996; Van Marrewijk et al., 1996), but differences in potency were documented for each AKH in L. migratoria, decreasing in the order AKH III > AKH I > AKH II (Vroemen et al., 1997). IP3, generated by activation of phospholipase C (PLC), plays a critical role in Ca2þ mobilization that leads to activation of protein kinase C (PKC) for a wide variety of neuropeptides (Berridge, 1993). The stronger effect of AKH I on IP3 combined with its weak effect on Ca2þ efflux relative to AKH II suggest that AKH I is the major lipid mobilizing hormone, particularly since Ca2þ is required for translocation of hormone sensitive lipases from the cytosol to lipid droplets to elicit lipolysis (Clark et al., 1991; Egan et al., 1992). The activation of glycogen phosphorylase by each of the AKHs is mediated by PLC activation, since a PLC inhibitor, U73122, dampened the response to the peptides, and the residual activity might be due to a cAMP pathway not influenced by IP3 (Vroemen et al., 1997). Elevation of IP3 levels by AKH is mediated by the G protein Gq, since a Gq antagonist dampened AKH I induced glycogen phosphorylase activity (Vroemen et al., 1998b). Stimulation of glycogen phosphorylase by cAMP is independent of extracellular Ca2þ (Van Marrewijk et al., 1993), and the elevation of cAMP by forskolin or db-cAMP did not affect the enhancement of InsPn levels by AKH I, so a direct linkage between

502 Hormones Controlling Homeostasis in Insects

Figure 2 Proposed model for the coupling of AKH signaling pathways in Locusta migratoria fat body cells for carbohydrate mobilization. R, receptor; G, G protein; PLC, phospholipase C; IP3, inositol-1,4,5-trisphosphate; AC, adenylate cyclase; GPh, glycogen phosphorylase. (Reprinted from Van der Horst,D.J., Vroemen, S.F., Van Marrewijk, W.J.A., 1997. Metabolism of stored reserves in insect fat body: hormonal signal transduction implicated in glycogen mobilization and biosynthesis of the lipophorin system. Comp. Biochem. Physiol. 117B, 463–474, with permission from Elsevier.)

the AKH receptor and PLC activation must exist that is not dependent on the elevation of cAMP. These data have led to a model describing AKH signaling in L. migratoria (Van der Horst et al., 1997) (Figure 2). The AKH receptor is coupled to at least two distinct G proteins, Gs and Gq, and the activation of both pathways results in a complete enhancement of glycogen phosphorylase activation by AKH. AKH induced Ca2þ influx is mediated by voltage independent channels, possibly calcium release activated channels, since La3þ, a universal Ca2þ channel blocker, prevents glycogen phosphorylase activation (Vroemen et al., 1998b). In P. americana, the effects of HrTH on phosphorylase activation in the fat body and trehalose release are dependent on the presence of extracellular Ca2þ (McClure and Steele, 1981; Orr et al., 1985; Steele and Paul, 1985), and the entry of Ca2þ into intact fat body (Steele and Paul, 1985), or fat body trophocytes (Steele and Ireland, 1999), is stimulated by HrTH. A key role for Ca2þ in glycogen breakdown is its requirement, together with calmodulin, for the activity of phosphorylase kinase, the activator of glycogen phosphorylase (Pallen and Steele, 1988). The mode of HrTH action in P. americana was examined in disaggregated trophocytes, the fat body cells responsible for trehalose synthesis (Steele and Ireland, 1994). This system was used to provide the ability to test the effect of peptides or varying conditions on a large number of samples to better correlate changes in a discrete population of responsive cells. It was demonstrated that in P. americana HrTH I and II stimulated IP3 formation with comparable potencies

(Steele et al., 2001). The HrTH stimulated formation of IP3 and the increase in intracellular Ca2þ concentration are probably involved in phosphorylase activation and trehalose efflux from trophocytes, since a PLC inhibitor blocked these effects. HrTH I and II increase Ca2þ levels in fat body trophocytes by the release of intracellular Ca2þ followed by the subsequent capacitative influx of extracellular Ca2þ (Sun and Steele, 2001). Capacitative Ca2þ entry is inhibited by activation of protein kinase C or inhibition of calmodulin (Sun and Steele, 2001). HrTH stimulates the activity of phospholipase A2 in trophocytes (Sun and Steele, 2002), which leads to the increased production of free fatty acids to provide an essential source of energy for trehalose synthesis (Ali and Steele, 1997). Phospholipase A2 activation and the stimulation of trehalose efflux are dependent on G proteins, protein kinase C and calmodulin, and occur following the increase in cytosolic Ca2þ mediated by HrTH (Sun et al., 2002; Sun and Steele, 2002). In P. americana, a CC extract elevated cAMP levels in the fat body, but this effect was not due to the action of HrTH (Orr et al., 1985). Transduction of the HrTH signal in B. discoidalis does not involve cyclic AMP, since HrTH does not elevate fat body cAMP levels (Park and Keeley, 1995), and elevating cAMP levels by IBMX or dbcAMP treatment does not lead to elevated trehalose synthesis (Lee and Keeley, 1994) or glycogen phosphorylase activity (Park and Keeley, 1995). Likewise, no effect of cAMP on trehalose production was observed in the cockroaches P. americana (Orr et al., 1985) or Blaptica dubia (Becker et al., 1998).

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The release of Ca2þ from intracellular stores by thimerosal or thapsigargin increased trehalose synthesis (Keeley and Hesson, 1995) and glycogen phosphorylase activation (Park and Keeley, 1996), even in the absence of extracellular Ca2þ. In contrast, stimulation of the influx of extracellular Ca2þ did not affect trehalose synthesis (Keeley and Hesson, 1995) and had only a modest effect on phosphorylase activation (Park and Keeley, 1996). However, a maximal response to hormone requires the presence of extracellular Ca2þ (Keeley and Hesson, 1995; Park and Keeley, 1996). Since HrTH was shown to increase IP3 levels in the fat body, it was suggested that HrTH leads to the release of intracellular Ca2þ followed by an influx of extracellular Ca2þ to achieve a maximal effect on the fat body (Park and Keeley, 1996). A similar dependence on extracellular and intracellular Ca2þ was demonstrated for HrTH mediated trehalose production in B. dubia (Becker et al., 1998). However, the HrTH mediated decrease in concentration of the metabolic signaling molecule, fructose 2,6-bisphosphate (Becker and Wegener, 1998), is largely dependent on Ca2þ entry, since this effect could be fully mimicked by the calcium ionophore A23187 (Becker et al., 1998). This decrease in fructose 2,6-bisphosphate (Becker and Wegener, 1998) is a key regulatory step towards the inhibition of glycolysis and directing the products of glycogen breakdown towards the production of trehalose that results from HrTH action (Wiens and Gilbert, 1967). The signal transduction pathway for the action of AKH on lipid mobilization in adult M. sexta involves both cAMP and Ca2þ (Arrese et al., 1999). The magnitude of diacylglycerol mobilization that was induced by Ca2þ mobilizing agents was similar to the response to the peptide, while treatments that increase cAMP levels did not induce a maximal response. For this reason, it was proposed that Ca2þ potentiates the action of cAMP. AKH induces an increase in PKA activity that precedes the activation of TAG lipase and release of diacylglycerol, further supporting the role for cAMP as a second messenger for AKH action in M. sexta. In the fruit beetle, Pachnoda sinuata, which mobilizes carbohydrate reserves and increases proline synthesis in response to AKH (Auerswald et al., 1998), the signal transduction pathway involves both cAMP and Ca2þ (Ga¨de and Auerswald, 2003). Injection of the endogenous AKH, Melme-CC (first isolated from M. melolontha), elevates cAMP levels in the fat body, and injection of a membrane-permeable cAMP analog or IBMX elevates proline levels in the hemolymph supporting the involvement of cAMP (Auerswald and Ga¨de, 2000). The induction of cAMP by AKH requires the presence of extracellular

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Ca2þ (Auerswald and Ga¨de, 2001a), and results in the rapid influx of Ca2þ into the fat body cell (Auerswald and Ga¨de, 2001b). The release of intracellular Ca2þ is required for the full stimulation of proline production by stimulating the influx of extracellular Ca2þ by capacitative Ca2þ entry (Auerswald and Ga¨de, 2001a). In contrast to the effect of cAMP on proline production, it is not involved in carbohydrate mobilization or glycogen phosphorylase activation (Auerswald and Ga¨de, 2000), which requires the presence of extracellular Ca2þ and the release of Ca2þ from intracellular stores (Auerswald and Ga¨de, 2001b). Injection of Melme-CC causes an increase in IP3 levels in the fat body, but coinjection with a PLC inhibitor, U73122, affects only the carbohydrate mobilizing activity, and not the hyperprolinemic effect of the peptide (Auerswald and Ga¨de, 2002). These data suggest that two separate pathways exist for AKH action in P. sinuata, and that the release of the IP3 dependent Ca2þ stores from the endoplasmic reticulum is not involved in proline production. 3.10.2.1.6. AKH receptor The M. sexta AKH receptor was characterized using tritium labeled Manse-AKH with a binding assay to larval fat body membrane preparations (Ziegler et al., 1995). Specific and saturable binding, which required the presence of Ca2þ, was detected with a Kd of 7  1010 M and was consistent with binding to one type of receptor. Smaller amounts of peptide are needed for a full response in a bioassay indicating that only a fraction of receptors must bind peptide for a maximal response. Specific binding to membrane preparations of M. sexta heart and muscle was not detected, but a low level of specific binding to the pterothoracic ganglion membrane preparations of adult M. sexta was observed (Ziegler et al., 1995), consistent with the action of AKH that was observed on central nervous system neurons (Milde et al., 1995). Two independent methods were used to identify the AKH receptor cDNA from D. melanogaster (Park et al., 2002; Staubli et al., 2002). In one study, the D. melanogaster receptor cDNA related to the gonadotropin releasing hormone (GnRH) receptor (AF077299) (Hauser et al., 1998) was expressed in a CHO cell line expressing the promiscuous G protein G16 (CHO/G16) and aequorin, and Drome-AKH was identified as the ligand present in a 3rd instar larval extract that induced a bioluminescent response (Staubli et al., 2002). A dose–response curve showed that the response induced by synthetic Drome-AKH had an EC50 of 8  1010 M, and all other D. melanogaster peptides tested were inactive. The D. melanogaster AKH receptor was independently identified by injecting in vitro synthesized

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p0215

RNA encoding the GPCR from the vasopressin receptor group (AF522194) into Xenopus oocytes and measuring the inward current in response to application of AKH (Park et al., 2002). Oocytes responded specifically to Drome-AKH with an EC50 of 3  1010 M. Xenopus oocytes expressing a second GPCR (AF522188) reacted with AKH, but with a much lower potency (EC50 ¼ 2.4  108 M). AF522188 also reacted with CCAP with an EC50 of 1.2  108 M suggesting that this GPCR might interact with multiple ligands. This suggestion is supported by the overlap in the functions of AKH and CCAP, since both peptides possess cardioacceleratory activity (Scarborough et al., 1984; Witten et al., 1984; Tublitz and Truman, 1985). The D. melanogaster AKH receptors independently identified are derived from the same gene, and differ only by a small deletion of 8 bp in AF522194 causing a frameshift in its predicted sequence at the C-terminus, and several point mutations, only one of which results in an amino acid substitution (Park et al., 2002). These differences did not affect the potency and specificity of their interactions with Drome-AKH. The characterization of an AKH receptor cDNA in one species leads towards the development of strategies to identify the AKH receptor in a wide variety of insect species (Staubli et al., 2002). Primers were designed to conserved regions of the D. melanogaster AKH receptor and a second receptor, that for D. melanogaster corazonin (Park et al., 2002), to clone the AKH receptor from Bombyx mori (Staubli et al., 2002). Aequorin expressing CHO/G16 cells transfected with the B. mori AKH receptor cDNA were activated by Helze-HrTH (Jaffe et al., 1988), a peptide not yet identified in B. mori, with an EC50 of 3  1010 M. The known AKH from B. mori (Ishibashi et al., 1992), identical to Manse-AKH and Helze-AKH, activated the B. mori AKH receptor at a lower affinity, suggesting that B. mori possesses a second AKH that is more related to Helze-HrTH. The presence of multiple AKH peptides in many insect species (Ga¨de et al., 1997), the differing rank order of potencies of these AKHs depending on the assay used (Goldsworthy et al., 1986b), and the age related changes in sensitivity to AKHs (Mwangi and Goldsworthy, 1977a; Ziegler, 1984; Woodring et al., 2002) raise the question of whether AKH(s) interacts with multiple receptors. The presence of multiple AKH receptors in some insect species has been suggested based on biphasic responses to peptide analogs (Ga¨de and Hayes, 1995). In P. sinuata, AKH probably acts through two receptors, since peptide analogs possess differential effects on carbohydrate mobilization and proline synthesis (Auerswald and

Ga¨de, 1999), and these biochemical effects are independently mediated by different signal transduction pathways (Auerswald and Ga¨de, 2000, 2002). However, evidence in M. sexta is consistent with the presence of one type of receptor responsible for the adipokinetic response, since analogs that are partial agonists are capable of reducing the response to the endogenous peptide (Fox and Reynolds, 1991a), and none of the AKH analogs tested exhibited a biphasic response, which would provide evidence for multiple receptors (Ziegler et al., 1998). Receptor desensitization, the increase in refractoriness in response to sustained exposure to the ligand, is mediated by phosphorylation of the agonist-bound receptor by G protein-coupled receptor kinases (Perry and Lefkowitz, 2002). Preincubation of L. migratoria fat body with any of the three Locmi-AKHs renders the tissue insensitive to each of the AKHs (Vroemen et al., 1998a). While this result might imply that all three AKHs share the same receptor, an alternative interpretation is that each receptor recognizes all three AKHs with differing affinities, and that prolonged preincubation with a high dose of one peptide could desensitize all receptors. 3.10.2.1.7. Effects on gene expression Flight activity is the only natural stimulus known for AKH release, but neither AKH mRNA levels nor prohormone synthesis were altered by flight activity (Harthoorn et al., 2001). Therefore, the availability of releasable AKH is dependent on its continuous biosynthesis. The first example of the regulation of HrTH gene expression by nutritional status was obtained in B. discoidalis (Lewis et al., 1998). It was demonstrated that starvation induced a twofold increase in HrTH mRNA levels in the CC, which returned to essentially normal levels after 2 days of refeeding. This starvation induced increase in HrTH mRNA level is accompanied by a similar increase in HrTH synthesis (Sowa et al., 1996). The downregulation of HrTH mRNA levels by feeding is not simply a response to elevated hemolymph carbohydrate levels or a neural response to feeding, but is more likely due to the consumption of a complex of nutrients (Lewis et al., 1998). The first example of an effect on downstream gene expression in response to an insect neuropeptide (HrTH) was demonstrated in the fat body of B. discoidalis (Bradfield et al., 1991). The mRNA level of a cytochrome P450 (P4504C1) was increased directly in response to physiological levels of HrTH, and was also upregulated by starvation in a CC-dependent manner (Bradfield et al., 1991; Lu

Hormones Controlling Homeostasis in Insects

et al., 1995, 1996). It was suggested that P4504C1 is involved in mobilization of fat body resources in response to starvation either by the control of fatty acid oxidation to provide energy for the conversion of glycogen to trehalose in response to the release of HrTH (Bradfield et al., 1991), or by providing an alternative route for carbohydrate synthesis during exposure to HrTH or starvation (Lu et al., 1996). 3.10.2.2. A Counter-Regulatory Hormone for AKH? p0245

Despite extensive research, until recently there was no evidence for a counter-regulatory hormone for the AKH family. Bombyxin is an insect homolog of insulin, both in structural properties (Kawakami et al., 1989) and receptor architecture and properties (Fullbright et al., 1997). However, treatment of the natural host with bombyxin results in an activation of glycogen phosphorylase and a decrease in glycogen stores (Satake et al., 1997), the opposite of the effects of insulin in vertebrates. An important role of insulin in vertebrates is lowering blood glucose, to not only maximize the utility of this expensive fuel for anaerobic metabolism, but to prevent damage to body proteins via formation of aldimines from the less abundant, but reactive, aldose form of glucose. Aldimines can rearrange to ketimines, which cannot hydrolyze like aldimines and thus permanently alkylate proteins with adverse effects. Because insect hemolymph contains the nonreducing sugar trehalose, this function of insulin would be of no importance to insects. Clynen et al. (2003a) used proteomic techniques to identify a small peptide, QSDLFLLSPK, from the GCC of L. migratoria; the mass spectral sequence was ambiguous, because this technique is unable to distinguish Leu from Ile, or Gln from Lys. However, a basic local alignment search tool (BLAST) search revealed a 100% match to a fragment of the locust insulin related peptide (Lagueux et al., 1990). This fragment is termed a ‘‘co-peptide’’ which is immediately downstream of the signal peptide in the prepro-insulin related peptide of L. migratoria. The co-peptide is just upstream of the B-chain, which is followed by the C-peptide, and then the A-chain. Each peptide is separated by dibasic amino acid cleavage sites. In the mature insulin related peptide, the A- and B-chains are joined by two disulfide bonds, with an additional, internal disulfide bond in the A-chain. The C-peptide is not known to have a biological role in vertebrates, and the co-peptide might be presumed to have no activity. Synthesis of the L. migratoria co-peptide and bioassay revealed that it inhibits glycogen phosphorylase in the fat body, an effect expected of an insulin-like hormone.

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However, it has no effect on hemolymph lipid levels. Bombyxin, the first insulin related peptide discovered in insects, actually lowers storage carbohydrates in Bombyx mori, a paradoxical effect for a homolog of insulin (Satake et al., 1997). The co-peptide certainly merits more detailed physiological study. Of some concern is the generality of these results: examination of the precursors of insulin related peptides in B. mori, A. gambiae, and D. melanogaster revealed that none of them contain the corresponding co-peptide; the B-chain in each begins immediately after the signal peptide. In the horsefly Tabanus atratus, two peptides with AKH activity have been identified: Tabat-AKH and Tabat-HoTH (Jaffe et al., 1989) (Figure 1). While Tabat-HoTH, a decapeptide, was claimed to have hypotrehalosemic activity (that expected for an insulinlike peptide rather than a member of the AKH/HrTH family), examination of Figure 5b in Jaffe et al. (1989) reveals that the curves showing dose-dependent differences between hemolymph sugars in control versus Tabat-HoTH treated larvae are but a trend and have overlapping standard deviations; no statistical tests were applied. Thus, the claims for hypotrehalosemic activity are unsupported by the published data.

3.10.3. Hormonal Control of Water and Electrolyte Homeostasis Insects are small and hence have a high surface area to volume ratio, making them vulnerable to desiccation. Nevertheless, insects have evolved to be the most populous and diverse of all terrestrial animals. Insects have cuticles covered with water-impervious hydrocarbons to minimize water loss, and employ a variety of other behavioral, morphological, and physiological adaptations to keep water loss to a minimum, particularly in species that occupy very dry habitats. To maintain fluid homeostasis, water gained from the diet, metabolism and, in some species, from the atmosphere, must equal water lost by evaporation, respiration, and excretion. Evaporative and respiratory losses vary with environmental factors and the state of hydration, but the major site of regulation is the excretory system, and water loss via this route can change dramatically depending on the physiological status of the insect. Excretory water loss is determined by the rate at which fluid enters the hindgut from the midgut and Malpighian tubules (MT), and the rate of reabsorption therein. Fluid and ion homeostasis are under endocrine and possibly neural control, which allows the insect to regulate hemolymph volume and composition while permitting nitrogenous waste, toxic substances, and excess ions and/or water to be voided. The

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endocrine factors affecting the regulation are referred to as diuretic hormones (DHs) and antidiuretic hormones (ADHs) although this may not precisely describe their actions. Generally, DHs stimulate primary urine secretion by MTs, whereas ADHs increase fluid reabsorption from the hindgut, but there are exceptions to this. 3.10.3.1. Molecular Basis for Urine Excretion

Urine excretion in vertebrates is primarily driven by blood pressure, but there is a complex interplay between a number of hormones in the control of fluid and ion homeostasis. Aldosterone and vasopressin (antidiuretic hormone) are largely responsible for maintaining electrolyte and water balance via reabsorption, while atrial natriuretic factor and angiotensin II primarily control blood volume (Beyenbach, 1993). In contrast, insects have low pressure circulatory systems and primary urine isosmotic to the hemolymph is secreted by the MT.

Diuresis must therefore be driven by active ion transport, not by blood pressure. Diuretic hormones increase MT secretion by stimulating ion transport, whereas antidiuretic hormones either reduce MT secretion or stimulate fluid and vital solute reabsorption in the hindgut (Phillips, 1983) (Figure 3). A large literature exists on diuresis and its regulation in insects; early physiological studies have been reviewed comprehensively (Phillips, 1983; Spring, 1990). However, most early physiological studies on hormonal control of fluid homeostasis in insects utilized crude extracts of insect neuroendocrine tissues to affect target organs. It is now known that such studies dealt with a mixture of different active factors, so that unraveling mode of action of any single factor was impossible. A fundamental understanding of the control of fluid homeostasis requires that the controlling factors be identified, synthesized, and tested for their effects. Only availability of synthetic factors can allow a detailed

Figure 3 A generalized scheme of the classic view of the excretory process in insects. Malpighian tubule secretion is driven by the active transport of KCl and/or NaCl into the lumen, which draws water by osmosis. Other ions and metabolites enter the lumen by passive or active transporters. The primary urine enters the gut in most species at the midgut–hindgut junction and generally is directed posteriorly into the ileum and rectum where it is modified by isosmotic fluid reabsorption. The fluid entering the ileum and rectum is extensively modified by reabsorption of essential metabolites and fluid, which may be hypo- or hyperosmotic to the luminal contents. The driving force for ion transport is an apical electrogenic Cl pump, with cations entering passively, in contrast to the Malpighian tubules. Toxic wastes are retained in the hindgut lumen and are voided in the excreta, which can be strongly hypo- or hyperosmotic to hemolymph. (Modified with permission from Coast, G.M., Orchard, I., Phillips, J.E., Schooley, D.A., 2002a. Insect diuretic and antidiuretic hormones. Adv. Insect Physiol. 29, 279–409.)

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molecular understanding of the actions of factors controlling excretion and their physiological interplay, and whether prime control is due to diuresis or antidiuresis. There have been a number of reviews of the hormonal control of MT, and of hindgut function, focusing on identified hormones (Audsley et al., 1994; Phillips and Audsley, 1995; Coast, 1996, 1998; Phillips et al., 1998a; O’Donnell and Spring, 2000), and an extremely comprehensive review of both MT and hindgut function was published recently (Coast et al., 2002b). The thrust of this review is to focus on advances in excretory physiology that concentrate on the effects of pure, defined factors. An understanding of the molecular basis of basal urine production in the Insecta has emerged only during the last 15 years, and has been reviewed by Nicolson (1993) and Beyenbach (1995, 2003). Ion transport leading to basal secretion of urine by the MT is driven by an apical vacuolar-type Hþ-ATPase (V-ATPase), in parallel with an apical cation/Hþ antiporter (Grinstein and Wieczorek, 1994). In a yellow fever mosquito, Aedes aegypti, MT, this transporter is believed to be a 1 cation:1 Hþ antiporter

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(Beyenbach, 2001), while in M. sexta midgut it is believed to be a 1 cation:2 Hþ antiporter (Azuma et al., 1995). The activity of the V-ATPase is stimulated by treatment of tubules with cAMP or peptides elevating cAMP, and cGMP also stimulates its activity in D. melanogaster (O’Donnell et al., 1996). The precise mechanism of stimulation is unknown; there is no evidence for phosphorylation of the V-ATPase, which lacks consensus sites for protein kinase A phosphorylation (Wieczorek et al., 2000). A body of evidence suggests that activation of the M. sexta protein is related to association of the V1 and V0 subunits via a redox process (Gru¨ ber et al., 2001); formation of a disulfide bond between Cys254 and Cys532 of the catalytic A protein of the yeast V1 unit leads to reversible inactivation of the V-ATPase (Forgac, 2000), apparently via dissociation of the knob-like V1 unit from the stalk-like V0 unit. The V-ATPase generates an electrochemical gradient favoring the entry of protons from the lumen in exchange for Kþ and/or Naþ (Bertram et al., 1991; Maddrell and O’Donnell, 1992; Weltens et al., 1992) (Figure 4). The alkali cation-Hþ antiports of the hematophagous insect R. prolixus have a

Figure 4 A generalized model for ion transport by Malpighian tubule principal cells. Active cation transport is energized by an apical (luminal) membrane Hþ pump (V-ATPase), which generates a gradient for protons to return to the cell via parallel Kþ/Hþ and Naþ/Hþ antiports. Where there is a favorable electrochemical gradient, cations may enter principal cells by passive diffusion through ion channels in the basolateral membrane (notably Kþ, but also Naþ in Aedes aegypti). Other routes for cation uptake include primary (Naþ–Kþ–ATPase) and secondary (Naþ–Kþ–2Cl and Kþ–Cl cotransport) active transport. Naþ entry may also be coupled to the transport of organic solutes (not shown). Cl that enters the cells via cation-Cl cotransporters probably exits through channels in the apical membrane. Arrows with circles indicate primary (black circles) or secondary (gray circles) active transport, while arrows through cylinders represent membrane ion channels. (Modified with permission from Coast, G.M., Orchard, I., Phillips, J.E., Schooley, D.A., 2002a. Insect diuretic and antidiuretic hormones. Adv. Insect Physiol. 29, 279–409.)

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preference for Naþ over Kþ (Maddrell, 1978; Maddrell et al., 1993b), whereas there appears to be no such preference in the house cricket, A. domesticus (Coast et al., 1991). Alkali cations enter principal cells through ion channels in the basolateral membrane, if the electrochemical gradient permits, or by secondary (e.g., cation/Cl cotransport) transport. The basolateral primary (Naþ–Kþ–ATPase) active transporter has long been proposed to be important in MT transport processes, yet its known stoichiometry in vertebrate systems (3 Naþ exported per 2 Kþ imported) is one opposing the observed direction of cation transport in MT. In fact, serotonin has been reported to inhibit the activity of the Naþ–Kþ–ATPase in MT of Rhodnius prolixus, with this inhibition claimed to be responsible for some of the diuretic activity of this amine (Grieco and Lopes, 1997). However, indirect evidence suggests it may not play a significant role in R. prolixus, because ouabain, a potent inhibitor of this transporter, has no effect on fluid secretion (Ianowski et al., 2002). The Naþ–Kþ–ATPase does not appear to be of functional importance in MT of A. aegypti (Weng et al., 2003) or in M. sexta (E. Chidembo and D.A. Schooley, unpublished data), as judged by a lack of vanadate or ouabain sensitivity of ATP hydrolase activity. In contrast, in T. molitor the Naþ–Kþ– ATPase seems to play an important role in maintenance of cell Kþ concentration, and ouabain is an irreversible inhibitor of MT fluid secretion (Wiehart et al., 2003). These species differences may be explained by the nature of cations secreted in primary urine; in the cryptonephric tubules of T. molitor, activities of Kþ exceeding 3 mol l1 have been reported (O’Donnell and Machin, 1991), whereas A. aegypti secrete large amounts of Naþ (Hegarty et al., 1991). Species or order differences in the relative activity of the Naþ–Kþ–ATPase may also underlie the high levels of hemolymph Kþ versus Naþ in many Lepidoptera, Coleoptera, and Hymenoptera (see Sutcliffe, 1963 and Section 3.10.3.12.1): a low activity of Naþ–Kþ–ATPase in epithelial tissues would remove the driving force for keeping blood Kþ levels low versus Naþ in hemolymph of most insect orders. In general, the electrochemical gradient over the epithelium favors Cl diffusion into the lumen through a shunt pathway. The Cl-selective shunt described by Pannabecker et al. (1993) in MT of A. aegypti does not appear to be in the principal cells, and is either paracellular or through a second cell type, the stellate cells, which form thin (3–5 mm deep) windows between the hemolymph and tubule lumen. Not all species have stellate cells; recently,

they have been shown to have a different embryonic origin than principal cells (Denholm et al., 2003). Agents elevating intracellular Ca2þ increase Cl conductance, but do not stimulate the V-ATPase. 3.10.3.2. A Multiplicity of Peptides Regulate Diuresis and Antidiuresis in Insects

A number of ‘‘model species’’ have been adopted by different investigators for studying diuresis and antidiuresis. The classic work of Ramsay (1954) was performed with a stick insect, Carausius morosus. A great deal of pioneering work on diuresis and its control has been done with the hematophagous hemipteran R. prolixus (Maddrell, 1963) and D. melanogaster has been proposed as a useful model for studying diuresis in insects (Dow et al., 1998; MacPherson et al., 2001). The Beyenbach group has studied ion transport mechanisms in A. aegypti for decades (Beyenbach, 2003). Locusts have been the preferred model for studying the role of the hindgut in fluid reabsorption, largely through the work of the Phillips group (Phillips and Audsley, 1995). The first hint that more than two different classes of peptides acted on MT was supplied by the Beyenbach group’s studies of regulation of A. aegypti tubules by nervous system extracts separated on HPLC (Petzel et al., 1985). The first undisputed identification of an insect DH was the identification from M. sexta of a 41 amino acid peptide with high sequence similarity to the sauvagine/urotensin/ urocortin/corticotropin releasing factor (CRF) family (Kataoka et al., 1989b). To date, 13 ‘‘CRF-like’’ DH structures have been identified by isolation and Edman degradation; four were identified by BLAST searches of the D. melanogaster, A. gambiae, and B. mori genomes (Figure 5). Somewhat later, the leucokinins (myotropic peptides originally isolated based on their ability to cause contractions of the cockroach hindgut) were also shown to have potent diuretic effects (Hayes et al., 1989). Currently, 33 different myokinin sequences are known from nine species of insects. In the last few years, it has become evident that a ‘‘cocktail’’ of neuropeptides are implicated in the regulation of MT secretion (Coast et al., 2002a; Skaer et al., 2002). The majority of these peptides have diuretic activity, stimulating fluid secretion. Others have antidiuretic activity and reduce the rate of secretion. From the mid-90s onwards, peptides belonging to four families in addition to the CRF-like DHs and myokinins have been shown to influence tubule secretion: the calcitonin-like (CT-like) DHs, CAP2b-like, tachykinin

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related peptides, and two antidiuretic factors, ADFa and ADFb. The greatest amount of research has been carried out on the first two families discovered, i.e., the CRF-like and myokinin families; these appear to be ubiquitous among insects, although there is considerable interspecific variation in activity. For example, the CRF-like DH of A. domesticus stimulates maximal secretion, and is about threefold more active than the native myokinins (Coast et al., 1992). Alternatively, in D. melanogaster and in M. domestica (Holman et al., 1999), the CRF-like DHs have poor activity compared with the native kinin. The situation is even more extreme in R. prolixus, where CRF-like peptides can elicit maximal secretion and myokinins appear to be inactive (Te Brugge et al., 2002). CT-like, CAP2b-like, and tachykinin related peptides may also be of widespread occurrence in insects, but have been investigated less extensively than the CRF-like DHs and myokinins. Indeed, to date, CT-like peptides have only been identified in three species, Diploptera punctata, D. melanogaster, and Formica polyctena (a partial sequence), and three in the genomes of A. gambiae, B. mori, and Apis mellifera, but these represent very diverse orders (Dictyoptera, Diptera, Lepidoptera and Hymenoptera, respectively). There is immunological evidence for the existence of these factors in six other insect species representing the Orthoptera, Coleoptera, Hemiptera, and Lepidoptera (see Section 3.10.3.5). Even less is known of the antidiuretic factors controlling MT secretion since these have only been described in T. molitor. Intriguingly, peptides that have diuretic activity in one species have been shown to have antidiuretic activity in another. Manse-CAP2b is a diuretic in Diptera (D. melanogaster and M. domestica), but an antidiuretic in R. prolixus and T. molitor. CAP2b has no effect on secretion by MT of M. sexta at 1 mmol l1 (Skaer et al., 2002), the latter organism being the host from which it was identified. Tenmo-ADFb is antidiuretic in T. molitor and A. aegypti, but a diuretic in A. domesticus (G.M. Coast, unpublished data). Even when comparing quite similar species, peptide activity can vary considerably, as for CAP2b in D. melanogaster and M. domestica. In D. melanogaster, CAP2b acts on principal cells to activate a calcium-calmodulin dependent NO synthase, and the rise in NO levels increases cGMP production by a soluble guanylate cyclase. Diuresis results from the cGMP dependent stimulation of an apical membrane V-type ATPase that powers fluid secretion (Davies et al., 1995). Conversely, CAP2b has no effect on cGMP levels in M. domestica tubules or on the activity of the V-type ATPase. Instead, diuresis

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results from the opening of a passive chloride shunt pathway, which has previously been attributed to myokinin stimulation (C.S. Garside and G.M. Coast, unpublished data). 3.10.3.3. CRF-Like DH

The first insect DH identified was Manse-DH, a 41 residue peptide isolated from 10 000 heads of pharate adult M. sexta using an in vivo assay (Kataoka et al., 1989b). This assay for monitoring the isolation utilized beheaded, newly eclosed adults of Pieris rapae, similar to a technique described by Nicolson for Pieris brassicae (Nicolson, 1976). Manse-DH has 41% sequence identity with sauvagine, a peptide isolated from skin of the frog Phyllomedusa sauvagei (Montecucchi and Henschen, 1981) and believed for some time to be the amphibian form of CRF. The direct sequence identity with CRF is somewhat lower. Subsequently a smaller peptide, Manse-DPII, was isolated from dissected neuroendocrine cells of brains of M. sexta. Again, an in vivo bioassay was used to monitor the isolation using adults of M. sexta. Due to the extremely small amount of tissue extracted in this ‘‘surgical purification’’ method, the peptide was obtained in a pure state after a single step of reversed-phase liquid chromatography (RPLC) (Blackburn et al., 1991). However, the amount of peptide isolated was so small that only a partial sequence was determined by Edman degradation; complete structure proof required mass spectral sequencing. As aligned in Figure 5, Manse-DPII has only nine residues identical with those in Manse-DH. Also in 1991, CRF-like DHs were isolated from A. domesticus and L. migratoria; the former was isolated based on its ability to increase levels of cAMP in MT of A. domesticus maintained in vitro (Kay et al., 1991a). This 46 amino acid peptide, Achdo-DP, is clearly a third member of the ‘‘CRF-like’’ DH. Two publications reported the sequence of the same DH from L. migratoria within a few weeks of each other; both utilized purifications by multiple steps of RPLC. Lehmberg et al. required only three RPLC purification steps to isolate this peptide to homogeneity from 4000 dissected locust brains using a direct enzyme-linked immunosorbent assay (ELISA) with antibodies raised against Manse-DH (Lehmberg et al., 1991). Kay and colleagues utilized whole heads of L. migratoria to isolate Locmi-DH, monitoring the four-step RPLC isolation procedure with the in vitro assay for cAMP production with MTs of the locust (Kay et al., 1991b). In 1992 Kay et al. identified from P. americana a 46 residue DH, which differs from Locmi-DH at 13 residues, and from Achdo-DP at 18 residues. MT of

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L. migratoria were used to monitor the isolation of this peptide, because of extreme variability between cockroach tubules (G.M. Coast, unpublished data). Much later, Furuya et al. (2000b) isolated a 46 residue DH from the Pacific beetle roach, Diploptera punctata, which still differs from Peram-DH at nine residues. Another 46 residue DH from Zootermopsis nevadensis (Baldwin et al., 2001), a dampwood termite, differs from PeramDH at only three residues, and thus is ranked phylogenetically between the two cockroach DHs by the sequence alignment program utilized, Clustal W version 1.8 (Figure 5). It is curious that these 46 residue DHs, while being relatively more related to one another than the other CRF-like DH, still vary highly in sequence. An interesting observation with Z. nevadensis is that this species contains another DH that elevates cAMP in tubules of M. sexta, the species used to assay the isolation of Zoone-DH. This second factor is more basic, but less hydrophobic, than Zoone-DH; two attempts to isolate it were unsuccessful, because of its lability in the acidic solvents used for extraction (Baldwin et al., 2001). The first, and to date only, case of two species having an identical CRF-like DH sequence was the identification of Musdo-DP from M. domestica and Stomoxys calcitrans (Clottens et al., 1994). The assay to monitor the purification scheme was

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based on elevation of cAMP in MT of M. sexta maintained in vitro. This 44 amino acid peptide was accompanied by another, more hydrophobic active factor in S. calcitrans. The existence of this unknown factor has been reported (see Schooley, 1994), but, unfortunately, no details of its properties were ever published. Audsley et al. (1995) advocated the use of M. sexta MT for a general heterologous bioassay for isolation of DH from other species, because these MT proved to cross-react more promiscuously with all CRF-like DH tested than MT of L. migratoria or A. domesticus. A DH was isolated from the salt water mosquito, Culex salinarius, the isolation of which has never been reported, although its sequence and biological properties have been published (Clark et al., 1998a). This 42 residue peptide differs from the Musdo-DH sequence at nine positions. Two more dipteran CRFlike DHs have been identified from the genome of D. melanogaster (Cabrero et al., 2002) and A. gambiae. Drome-DH44 differs from Musdo-DH at only one residue: the substitution of a Ser at position 31 for a Thr in Musdo-DH. In contrast, Anoga-DH44 differs from Musdo-DH by seven residues. All dipteran DHs isolated to date are two residues (in one case four residues) shorter than the 46-mers isolated from the Orthoptera, Dictyoptera, and a single isopteran.

Figure 5 The CRF-like DHs and the CRF superfamily of peptides – the blue colored residues emphasize the similarities among the CRF-like DHs. Sauvagine, urotensin I, and urocortin 1 are paralogs of CRF. The CRF-like DHs are shown using the Swiss Prot species abbreviations (defined in the text except for Bommo (Bombyx mori, whose genome sequence appeared while the article was in press)); wsUrot I, Catostomus commersoni urotensin I; Xenla, Xenopus laevis. The four identical CRFs are from human (h), rat (r), goat (c), and horse (e). Two residues (shown in red) are conserved in the lepidopteran DHs and the CRF superfamily. The unique C-termini of the Tenebrio molitor DH are shown in green.

Hormones Controlling Homeostasis in Insects

An unusual CRF-like DH was identified in T. molitor (Furuya et al., 1995); its isolation was monitored by the elevation of cAMP from MT of this species maintained in vitro. This 37 residue DH, Tenmo-DH37, was isolated from 8400 pupal heads of T. molitor, and was accompanied by a less abundant, more hydrophobic factor isolated in insufficient quantity to sequence. Only later was the structure of this second factor reported, after extraction of an additional 20 000 heads, combined with the active fractions from identification of Tenmo-DH37. The peptide isolated, Tenmo-DH47, is the largest CRF-like DH identified to date (Furuya et al., 1998) at 47 residues, and is much less abundant than its small congener. Structurally, both peptides differ from all other CRF-like DH in having their C-terminal residue existing in the free carboxylate form rather than being amidated. In addition, Tenmo-DH37 has a one-residue ‘‘extension’’ of its sequence: all sequences from organisms other than T. molitor have a C-terminal Ile-amide or Valamide, whereas Tenmo-DH37 has a Leu-Asn-OH C-terminus, and Tenmo-DH47 has a Leu-OH, lacking the Asn extension of its smaller congener. A physiological consequence of this structural difference is that Tenmo-DH37 is without any detectable biological activity on MT of M. sexta: its activity is 104 lower than Manse-DH in this assay. This is consistent with data on the free acid form of Manse-DH which is about 1000-fold less active than the natural, amidated form in the P. rapae assay (Kataoka et al., 1989b). The more abundant, smaller Tenmo-DH37 was found to be about 600-fold more potent than the larger peptide in an in vitro assay measuring cAMP production with adult MT (Furuya et al., 1998), and about 200-fold more potent in a Ramsay fluid secretion assay with larval MT (Wiehart et al., 2002). However, Manse-DH is only 17-fold less active than Tenmo-DH37 in the cAMP assay with adult MT (Furuya et al., 1995). The two latter DHs have only 15 residues in common in the alignment in Figure 5, with Tenmo-DH37 being more similar to Manse-DPII, and Tenmo-DH47 being more similar to the ‘‘longer’’ CRF-like DH. Two DHs were isolated from the white-lined Sphinx, Hyles lineata, a close relative of M. sexta (Furuya et al., 2000a). The larger of these, HylliDH41, differs from Manse-DH only at residue 27, which is Gln rather than His (Manse-DH). Similarly, the shorter DH, Hylli-DH30, differs from Manse-DPII only at residue 9, by a conservative substitution (Glu in the H. lineata peptide for Asp in Manse-DPII). Thus, the sequences of two DHs from each of three species are currently known, and

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it is apparent that these form a group of paralogous sequences. It is fascinating to contrast the CRF-like DH with the CRF superfamily of peptides in vertebrates. In 1991, two CRF sequences were identified by cDNA cloning in the white sucker Catostomus commersoni. The sucker CRFs differ from the sequences of human, horse, rat, and dog CRF (all identical) at only one and two residues (Morley et al., 1991), yet are only 54% identical with urotensin, the fish equivalent of sauvagine. The following year, a similar situation was discovered with Xenopus laevis, which has two forms of CRF differing from the human sequence at only 2 and 3 residues (Stenzel-Poore et al., 1992). Consequently a sauvagine (or urotensin)-like peptide was sought and found in rat brain: urocortin 1, a vertebrate ortholog of urotensin and sauvagine (Vaughan et al., 1995). Thus, the CRF superfamily in vertebrates consists of two paralogous subfamilies of sequences, a situation mirrored in the CRF-like DH, although the sequences are very highly conserved in vertebrates in contrast with those known from insects. Examination of the sequences of the 15 known CRF-like DHs (Figure 5) reveals only four amino acids that are completely conserved in this family. Two of these are a Ser and a Leu, but they are separated by seven amino acids. The others are an Asn and a Leu in the C-terminal domain, separated by three amino acids. The Leu is separated from the C-terminal residue by two amino acids (three in Tenmo-DH37). Consequently, the prospects for using degenerate probes to attempt to isolate additional new members of this family from cDNA or mRNA are very poor. Curiously, in the CRF superfamily of peptides, again only four residues are completely conserved; one is the Ser also conserved in the CRF-like DH, and another the Asn near the C-terminus. The other two totally conserved residues in the CRF superfamily differ from those in the CRF-like DH. Despite the exceptional sequence conservation of the various forms of CRF, the paralogs sauvagine, urotensin, and urocortins diverge considerably (Hauger et al., 2003). Aside from the genome sequences for DromeDH44, Boomo-DH41, Bommo-DH34, and AnogaDH44, to date only one gene sequence encoding a CRF-like DH has been determined by cloning; that is for Manse-DH (Digan et al., 1992). The prepropeptide has a 19 residue signal peptide for cellular secretion, followed by a 61 residue propeptide terminating in Lys-Arg (the cleavage site for a Kex-2 like processing enzyme), then the 41 residues of Manse-DH, followed by a Gly-Lys-Arg plus 15 additional amino acids of the propeptide. The

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Lys-Arg processing site results in a C-terminal Gly, which provides the amidation on the C-terminal Ile following action of peptidylglycine amidating monooxygenase (Prigge et al., 2000) on the Gly residue. Another aspect of the sequence alignment is worth noting. A Pro residue is situated either exactly between the totally conserved Ser and Leu residues in the N-terminal domain, or one residue closer to the Ser in two of the three ‘‘short’’ CRF-like DHs. The residue just upstream of this Pro is Asn in 11 of the 15 CRF-like DH, Ala in two, and Leu in two. The Asp-Pro bond, not observed in any DH isolated to date, is exceptionally labile to acid on standing in solution, and the Asn codon requires but a single base change to give Asp. The inability to isolate the second CRF-like DH from Z. nevadensis was probably due to the presence of an Asp-Pro bond (Baldwin et al., 2001); multiple attempts to isolate a CRF-like factor from heads of A. mellifera, assayed by its strong stimulation of MT of M. sexta, were totally unsuccessful because the activity would disappear after only a few steps (E. Lehmberg and D.A. Schooley, unpublished data). There are very few studies on structure–activity relationships with the CRF-related DHs because of their size and difficulty of solid-phase chemical synthesis. Studies with Manse-DH and Achdo-DP show that a region near the N-terminus is important for receptor activation while the remaining portion of the peptide is required for receptor binding. An analog Manse-DH(13–41), truncated by 12 residues at the N-terminus, binds Manse-DHR expressed in Sf9 cells with high affinity (IC50
2.8 nmol l1 versus 0.16 nmol l1 for the intact peptide) but does not stimulate cAMP production, whereas Manse-DH (3–41) has high receptor binding affinity and is a potent stimulant of adenylate cyclase (Reagan, 1995a). Manse-DH(21–41) and Manse-DH(26–41) have binding affinities reduced 100-fold and 1000fold, respectively, while Manse-DH(31–41) has no binding activity at 1 mmol l1 (Reagan et al., 1993). Deletion of the first four residues from Manse-DH decreases activity, while deletion of the next residue essentially abolishes it (P. Dey and D.A. Schooley, unpublished data). Taken together with the results of Reagan (1995a), this suggests that the receptor activating domain of Manse-DH lies very close to the N-terminus. Studies with N-terminal truncated analogs of Achdo-DP in a cricket tubule fluid secretion assay confirm the importance of the N-terminus for receptor activation (Coast et al., 1994). The activities of Achdo-DP(6–46) and Achdo-DP(7–46) are essentially indistinguishable from intact Achdo-DP, but the activity of Achdo-DP(11–46) is reduced by

60% and Achdo-DP(23–46) is inactive. These data could suggest that the domain necessary for receptor activation is farther from the N-terminus in AchdoDP than in Manse-DH. However, results for peptides with low activity in a series of deletion peptides must be interpreted with caution; such peptides are made by removing peptide synthesis resin at various stages of the synthesis, which proceeds from the C- to the N-terminus. Usually all synthetic deletion peptides are purified using the same preparative RPLC column; traces of biological activity from a fulllength, or nearly so, peptide will bleed out of a column essentially permanently (D.A. Schooley, unpublished data). Thus, the shortest peptides must be purified first on a new column. Intact Met residues may be important for activity; one of the Met residues (at position 1, 3, or 13) in Locmi-DH can become oxidized, with a resultant loss of activity (I. Kay and G.M. Coast, unpublished data). This may explain why [Nle2,11]-Manse-DH (with Nle replacing Met2 and Met11) maximally stimulates cricket tubule secretion, whereas Manse-DH gives only a 60% response (Coast et al., 1992). In addition, Locmi-DH(1–23) and Locmi-DH(24–46) are inactive whether tested separately or together (Nittoli et al., 1999) (G.M. Coast, unpublished data); thus, the binding and activation domains must be joined together to have a biologically active peptide. Manse-DH with a free C-terminus (acid) stimulates cAMP production by adult M. sexta MT (Audsley et al., 1995) and posteclosion diuresis in decapitated newly emerged P. rapae (Kataoka et al., 1989b), but in both assays it was 1000-fold less potent than Manse-DH. Reagan et al. (1993) were unable to detect any binding of this ligand to the Manse-DHR, which they attributed to rapid degradation of the nonblocked peptide (presumably by carboxylpeptidases) in the receptor binding assay. 3.10.3.4. Diuretic and Myotropic Peptides: the Kinins

Insect kinins (originally called myokinins) were isolated from whole head extracts of the Madeira cockroach, Leucophaea maderae (leucokinins; Leuma-Ks) (Holman et al., 1986a, 1986b, 1987a, 1987b) based on their potent myotropic activity in a cockroach (L. maderae) hindgut assay (Holman et al., 1991). The eight leucokinins differ in myotropic potency. They were later found to be potent stimulants of fluid secretion in A. aegypti, also causing a depolarization of the transepithelial potential (Hayes et al., 1989). Soon thereafter, five kinins (achetakinins; Achdo-Ks) were isolated from A. domesticus using the L. maderae hindgut assay; they were shown to also possess high diuretic activity on MT

Hormones Controlling Homeostasis in Insects

of A. domesticus (Coast et al., 1990). Similarly, locustakinin was isolated from 9000 brain–corpora cardiaca–corpora allata–subesophageal complexes of L. migratoria using the L. maderae hindgut assay (Schoofs et al., 1992). It is curious that only a single kinin, locustakinin (Locmi-K), was isolated from this species, whereas all other orthopterans and dictyopterans studied to date have 5–8 kinins. Using hindgut of P. americana, eight kinins were isolated from 800 corpora cardiaca–corpora allata of this species (Predel et al., 1997), five of which are unique sequences, while three occur in other species: Locmi-K, Leuma-K-7, and Leuma-K-8. Availability of endogenous kinins is crucial to understanding their role in any species. For example, muscakinin is about 106 more potent in M. domestica (Holman et al., 1999) than leucokinin I. The first kinins to be identified from a holometabolous species were those from A. aegypti, which were isolated using an ELISA rather than a functional assay (Veenstra, 1994). While these were termed ‘‘Aedes leucokinins 1–3,’’ this nomenclature suggests they may be the same sequence as those from L. maderae, but they are in fact unique. Here the Swiss-Prot-NCBI five-letter standards for species abbreviation, Aedae-K-1, Aedae-K-2, and AedaeK-3, as previously recommended is used (Coast et al., 2002a). Later, the sequence of the single cDNA encoding these kinins was determined, and their biological properties determined (Veenstra et al., 1997). While all three Aedae-K peptides are capable of depolarizing the mosquito MT at doses between 0.1 and 1 nmol l1, only Aedae-K-3 gave good stimulation of fluid secretion, with an EC50 near 10 nmol l1; Aedae-K-3 was appreciably less active and Aedae-K-2 was without effect on fluid secretion at 1 mmol l1. Three kinins were isolated from wild-collected salt marsh mosquitos, 94% of them C. salinarius, and the first kinin was christened ‘‘culekinin depolarizing peptide’’ (Hayes et al., 1994). They were isolated using the L. maderae hindgut assay, as well as an electrophysiological assay with tubules of A. aegypti. The sequence of only the heptamer Culsa-K-1 was published (Hayes et al., 1994). Some years later, the biological properties of Culsa-K-2 and Culsa-K-3, along with the sequences, were published by other workers (Cady and Hagedorn, 1999a, 1999b), but details of their isolation and identification remain unpublished. These peptides stimulate inositol trisphosphate (IP3) concentrations in MTof A. aegypti, with no effect on cAMP levels (Cady and Hagedorn, 1999b). Further, they stimulate in vivo urine production in adult female mosquitos (Cady and Hagedorn, 1999a). Using the

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sequences of the Aedae-K as a BLAST query of the A. gambiae genome (Holt et al., 2002), a single gene encoding three different kinins of 15, 10, and 21 residues was located: Anoga-K-1, -K-2, and -K-3. These peptides occur in the gene in the size order shown; the first and second are separated only by the amino acid residues Gly-Lys-Arg, as are the second and third (residues necessary for cleavage from the propeptide and formation of the amidated C-terminus). There are no reports of their synthesis and biological properties as yet. Holman et al. (1999) isolated a 15 residue kinin from another fly species, M. domestica, using an ELISA with an antibody raised against Leuma-K-1. Musdo-K strongly stimulates fluid secretion in M. domestica MT, whereas the hexapeptide AchdoK-5, identical with the six C-terminal residues of Musdo-K, is 1000-fold less potent in this assay. Thus, the longer sequence is crucial for full activity in M. domestica. Musdo-K also stimulates the housefly hindgut, possibly aiding in excretion. Later that year, Terhzaz et al. (1999) reported the isolation of a single 15 residue kinin from D. melanogaster, using a purification strategy based on that for the Aedae-K peptides. The sequence differs from Musdo-K at the second residue, a Thr in Musdo-K versus a Ser in Drome-K. A portion of the gene encoding a partial propeptide was determined from a BLAST search of sequenced bacterial artificial chromosomes of the then-incomplete D. melanogaster genome. Studies of the effects of Aedae-K on MT showed that it elevates intracellular Ca2þ, but not cAMP. A BLAST search of the now-complete genome of this species verifies that the pp gene encodes but a single kinin. The isolation of kinins from a lepidopteran insect, the corn earworm Helicoverpa zea, is of interest because only the first step utilized a functional assay. Blackburn et al. (1995) used
2000 abdominal ventral nerve cords (VNC) from adult moths as tissue source. Homogenized tissues were prepurified on a Sep-Pak C18, and then fractionated on RPLC. Fractions were assayed using a Ramsay assay with MT of adult M. sexta. Three fractions with diuretic activity coincided with peaks having strong absorbance at 280 nm, which gave second derivative UV spectra characteristic of Trp, one of three amino acids totally conserved in the kinins. Subsequent purification steps were monitored based on the diagnostic UV spectral behavior of Trp, rather than by using a functional assay. The peak eluting first was purified to homogeneity in one more RPLC step, and sequenced by Edman degradation (Helze-K-III; see Figure 6). The other two peptides, present in

514 Hormones Controlling Homeostasis in Insects

Figure 6 A multiple sequence alignment (Clustal W, V. 1.8) of the 33 known insect kinins from nine species. Boxed residues are identical. At the C-terminus, a five amino acid motif is highly conserved: FX1X2WGamide. X1 is N, S, Y, or H in all known sequences. X2 is usually S, but may be P or A. The first kinins were isolated from Leucophaea maderae (Leuma). For the definition of other species’ abbreviations, see text. Note most of the dipteran kinins cluster towards the center of the alignment. There is immunocytochemical evidence for the presence of kinins in over 15 species of insects. Interested parties are referred to a recent, massively comprehensive review (Coast et al., 2002a).

smaller quantity, required two additional steps for purification to homogeneity. The Trp residue could not be detected by Edman sequencing in these two kinins, which were additionally sequenced by tandem MS sequencing methods. The deduced sequences of all Helze-Ks were synthesized, and bioassayed on MT of M. sexta. No EC50 values were determined, only ‘‘threshold values’’: the lowest concentration at which a significant effect on fluid secretion can be measured. These were
7 pmol l1 for Helze-K-I, 0.6 pmol l1 for HelzeK-II, and 6 pmol l1 for Helze-K-III. None of these peptides had activity in vivo in H. zea, but they do not appear to have been tested in vitro in H. zea, or in vivo in M. sexta, so no valid comparison can be drawn. While kinins have not yet been identified in R. prolixus, partially purified extracts from neuroendocrine tissue of this species, which contain kinin-like immunoreactivity, do not stimulate tubule secretion, but are active on the hindgut (Te Brugge et al., 2002).

3.10.3.5. Calcitonin-Like DH

The first member of this family was identified together with the CRF-like Dippu-DH46 isolated from 4000 brains of D. punctata (Furuya et al., 2000b). During the first step of RPLC purification of extracts, only one fraction (corresponding to Dippu-DH46) stimulated cAMP production by MT of M. sexta, yet this fraction and a faster eluting fraction both elevated cAMP production by MT of S. americana. Because the M. sexta assay is easier, it was utilized to isolate Dippu-DH46. Isolation to homogeneity of the second factor was accomplished using MT of S. americana. Upon sequence analysis, it was immediately apparent that this 31 residue peptide was not a member of the CRF-like DH. A BLAST search revealed similarities to certain proteins, but no highly significant resemblance to any bioactive peptides. Nevertheless, a manual search through the catalog of a commercial supplier of peptides (Peninsula Laboratories) for the unusual GP-NH2 C-terminal sequence revealed an interesting similarity to

Hormones Controlling Homeostasis in Insects

p0370

f0040

calcitonin; in fact, only the calcitonin family has a Pro-NH2 at the C-terminus of all bioactive peptides listed in this catalog. In the alignment shown in Figure 7, only 6 of 31 residues of Dippu-DH31 are well conserved in the calcitonin family. However, chicken calcitonin is only 30-fold less potent in a fluid secretion assay than Dippu-DH31 using MT of D. punctata (Furuya et al., 2000b), suggesting this to be a genuine homology: one of the main actions of calcitonin is lowering blood Ca2þ by increasing Ca2þ excretion by the kidneys, in addition to increasing Ca2þ deposition in bone, which require active transport mechanisms. Interestingly, DippuDH31 and Dippu-DH46 have potent synergistic interaction in D. punctata (Furuya et al., 2000b). The actions of Dippu-DH31 in L. migratoria are more consistent with an elevation of intracellular Ca2þ in this species (Furuya et al., 2000b). These apparent differences in signaling are not surprising in light of the fact that a single, cloned calcitonin receptor has been shown to be capable of activating both the adenylate cyclase and phospholipase C signal transduction systems (Chabre et al., 1992; Force et al., 1992). To date, the only other characterized CT-like DH is that cloned ‘‘in silico’’ from the genome of D. melanogaster (gene Dh31 CG13094); this peptide, Drome-DH31, was synthesized and shown to be a potent diuretic in fruit fly tubules, and to act via adenylate cyclase (Coast et al., 2001). Another homolog of Dippu-DH31 exists in the genome of A. gambiae (BX038390) (Riehle et al., 2002), Anoga-DH31, but its biological function has not yet been studied. Moreover, with the recent availability of preliminary genome sequence data for B. mori and A. mellifera, genes encoding CT-like homologs exist in these two species (Bommo-DH31

Figure 7 The known calcitonin-like DHs are shown using the Swiss Prot species abbreviation convention: Forpo, Formica polyctena; other species abbreviations are given in text. In this figure, red is conserved in the calcitonin family; blue is conserved in Dippu-DH31 and homologs. Very recently (while this article was in press), the Bombyx mori (Bommo) genome because available. Limited genome sequence data also became available for Apis mellifera (Apime).

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and Apime-DH31). Using a direct ELISA with an affinity-purified Dippu-DH31 antibody, immunoreactivity was observed in head extracts of M. sexta, T. molitor, R. prolixus, A. domesticus, Blaptica dubia, and the Mormon cricket, Anabrus simplex (V.C. Lombardi, J.J. Hull, V. Te Brugge, and D.A. Schooley, unpublished data). At the time of writing, isolation of the M. sexta ortholog of Dippu-DH31 is imminent. Thus, it is likely that the CT-like DHs constitute another family of DHs of widespread occurrence within the Insecta. The partial sequence of a similar peptide isolated from the Belgian forest ant, Formica polyctena, was published in a Ph.D. thesis (Laenen, 1999). The isolation of the factor was monitored by observing its stimulation of MT writhing in L. migratoria: sequence analysis of this peptide was only possible out to 29 residues, but all are identical with the first 29 residues of DippuDH31 (see Figure 7). MS analysis revealed that the sequence was incomplete; the last two amino acids not detected must be different from those in Dippu-DH31 because the Mr is lower than that of the latter peptide (Laenen, 1999). Because the peptide lacked any diuretic activity on tubules of F. polyctena, from which it was isolated, it was not investigated further. Unfortunately, this discovery predated the publication of the sequence of DippuDH31; the sequence identity might have spurred completion of the sequence of the ant peptide despite its lack of diuretic activity in the host species. 3.10.3.6. CAP2b/Periviscerokinins

Manse-CAP2b was isolated from M. sexta as a cardioactive factor (Huesmann et al., 1995). Davies et al. (1995) found that it stimulates fluid secretion from MT of D. melanogaster by the unusual action (for a peptide hormone) of elevating production of nitric oxide, which in turn activates a soluble guanylate cyclase. This peptide was reported, based exclusively on the retention time of biological activity from D. melanogaster tissue extracts on RPLC, to be an endogenous peptide in D. melanogaster. However, a BLAST search of the D. melanogaster genome (Vanden Broeck, 2001) revealed the existence of three homologs of CAP2b with 8, 10, and 12 amino acid residues, having 6, 7, and 5 amino acid identities, respectively, to Manse-CAP2b (Figure 8). Garside and Coast (unpublished data) have studied the activity of Manse-CAP2b and Drome-capa-2 on MT of houseflies; in this species they are diuretic but signal transduction is not via NO or cGMP, but appears to result from an increase in intracellular Ca2þ. Predel et al. (1995) isolated a myotropin from perisympathetic organs of the cockroach P. americana christened periviscerokinin, which

516 Hormones Controlling Homeostasis in Insects

Figure 8 The Manduca sexta CAP2b family of peptides. All are amidated at the C-terminus (indicated with lower case a), and two (Manse-CAP2b and Anoga-CAP2b9 have pyroglutamic acid as the N-terminal residue (denoted by pQ, because Gln (Q) is the precursor of pGlu). At least one of the peptides with low sequence similarity in the C-terminal domain (FPRVa), PeramPVK-1 (from Periplaneta americana), has no diuretic activity, suggesting that the conserved C-terminal motif is crucial for activity.

seemed to be a peptide unrelated to other families of peptides. However, subsequently, this group (Predel et al., 1998) isolated a second peptide from this source, Peram-PVK-2, whose sequence was obviously related to not only Peram-PVK-1 but also to CAP2b. The CAP2b family now consists of 14 members, with the addition of three peptides from the cockroach L. maderae (Predel et al., 2000), one from the locust (Predel and Ga¨de, 2002), and three from the genome of A. gambiae (Riehle et al., 2002) (Figure 8). Recently, partial sequences for two new members of this family were identified using highly sensitive mass spectrometric assays (Clynen et al., 2003b) on single perisympathetic organs from two fly species, Neobelleria bullata and M. domestica. These sequences are incomplete because of the inability of MS analysis to distinguish between Ile and Leu in peptides, and because of lack of certain crucial diagnostic ions and are therefore not given in Figure 8. It is novel that this approach to isolation and identification does not require a bioassay. 3.10.3.7. Arginine Vasopressin-Like Insect Diuretic Hormone

Proux et al. (1982) found that a factor in the subesophageal ganglia of L. migratoria stimulated in vivo clearance of injected amaranth from the hemolymph of locusts, an established assay for diuresis in this species (Mordue, 1969), and obtained evidence that this material was identical to a factor that cross-reacts with arginine vasopressin (AVP). This factor was isolated from 51 000 dissected subesophageal and thoracic ganglia using RPLC techniques with monitoring of fractions with a radioimmunoassay for AVP. In the first step of isolation two factors were separated, both of which were

purified to homogeneity (Schooley et al., 1987). Injection of the purified factors into S. gregaria revealed that only the less abundant, slower eluting factor (F2) caused amaranth excretion. The faster eluting F1 and slower eluting F2 were found to have identical amino acid compositions and identical primary sequences: Cys-Leu-Ile-Thr-Asn-Cys-Pro-ArgGly-NH2. This peptide was synthesized in both the amidated and free acid forms, and after reduction and carboxymethylation was compared with the similarly modified forms of F1 and F2 prepared for sequence analysis. This comparison established that F1 was identical to the nonapeptide whose sequence is shown above. Analysis of both materials by size exclusion chromatography suggested that F2 was a dimer of F1. Two separate, difficult specific syntheses were performed to prepare pure F2 as parallel and antiparallel dimers (Proux et al., 1987). The synthetic antiparallel dimer proved to have identical retention properties on RPLC with F2 (Proux et al., 1987). Natural and synthetic F2 were shown to promote fluid secretion in an unusual fluid secretion assay in which a group of L. migratoria MT attached to the ampulla were removed, maintained in oxygenated saline, and the high combined flow of all tubules monitored (Proux et al., 1988). Later, this work was called into question by Coast et al. (1993), who reported that synthetic F2 had no stimulatory activity on a single L. migratoria MT in a conventional Ramsay assay, although Locmi-DH (a CRF-like peptide) had potent activity. Later, Montuenga et al. (1996) showed that there are endocrine cells in the ampulla of L. migratoria that contain not only Locmi-DH, but a second, as yet uncharacterized peptide, which is recognized by a Locmi-DH antiserum but is faster eluting on RPLC than the latter peptide. It is conceivable that the ‘‘AVP-like DH’’ may stimulate the release of Locmi-DH from these endocrine cells. The localization of AVP-like immunoreactivity has been determined in the locust (Thompson et al., 1991); the titer of AVP-like immunoreactivity changes in the hemolymph in response to unknown stimuli and neurons containing it innervate nonocular photoreceptors (Thompson and Bacon, 1991). Analysis by RPLC of AVP-like containing neurons showed them to contain not only the antiparallel dimer F2, but also the parallel dimer and the monomer F1 (Baines et al., 1995). Thus, there is strong evidence that the AVP-like factor is a bona fide neuropeptide, although its role remains controversial. 3.10.3.8. Other Peptides with Diuretic Activity

Skaer et al. (2002) tested three different tachykininrelated peptides (TRP) on MT of pharate adult

Hormones Controlling Homeostasis in Insects

p0400

M. sexta: locustatachykinin-1 from L. migratoria (Locmi-TK-1; GPSGFYGVRamide; Schoofs et al., 1993) and two TRPs from L. maderae (Leuma TRP-1, APSGFLGVRamide; and Leuma-TRP-4, APSGFMGMRamide; Muren and Na¨ ssel, 1996). Each TRP was tested at four concentrations from 1 nmol l1 to 1 mmol l1. Each TRP exhibited a dose dependent increase in the rate of fluid secretion, but a maximal response was reached only 30–40 min after application. While no EC50 values were measured, they appeared to be in the range of 10–100 nmol l1. Leuma-TRP-1 gave the highest maximal secretion of the three; at 1 mmol l1, it increased the rate of fluid secretion 2.83-fold compared to that of the control. In contrast to the CAPs and leucokinin, TRP effects on tubule secretion activity were not long lasting. Two cardioactive peptides of as yet unknown sequence, CAP1a and CAP1b (Tublitz et al., 1991), strongly stimulate M. sexta MT; there is a distinct probability they could be kinins. The very slow response (20–30 min delay) observed as a result of treatment of tubules with all agonists may be attributable to their use of 8–12 cm long pieces of a single MT (Skaer et al., 2002); M. sexta tubules tend to collapse on dissection and the delay in fluid secretion may result from the filling time of the tubule (G.M. Coast, unpublished data). Recently, Locmi-TK-1 has been reported to have an EC50 of 1.2 nmol l1 in L. migratoria, with an identical value reported for S. gregaria (Johard et al., 2003). The response observed was about 75% of that observed in response to treatment with LocmiDH; the latter was found to have synergistic activity with both Locmi-TK-1 and serotonin. In light of the response observed to this peptide in these two distantly related species (M. sexta versus locusts), further studies of the effects of these peptides will be of very great interest. Three fractions were isolated from a head extract of A. aegypti by RPLC based on their effect on the transepithelial potential (TEP) of isolated perfused MT (Petzel et al., 1985). Each fraction contained a pronase-sensitive peptide with Mr values estimated by gel filtration chromatography to be 2400 (fraction I), 2700 (fraction II), and 1860 Da (fraction III) (Petzel et al., 1986). Fraction I depolarized the TEP. While it has no effect on fluid secretion, it increases tritiated water (THO) loss and urine output from intact flies, possibly by inhibiting fluid uptake from the hindgut (Wheelock et al., 1988). Fraction II also depolarized the TEP, with a biphasic response; the TEP first depolarized, and then hyperpolarized. Fractions II and III both have diuretic activity and selectively stimulate secretion of NaCl-rich urine.

517

Fraction III was the more potent of these two; its diuretic and natriuretic activity were indistinguishable from that of exogenous cyclic AMP, although the latter only hyperpolarized the TEP (Petzel et al., 1985). Fraction III was named mosquito natriuretic peptide (MNP) and was shown subsequently to stimulate cyclic AMP production in isolated tubules (Petzel et al., 1987). Hegarty et al. (1991) found that dibutyryl cAMP stimulated Naþ secretion from A. aegypti MT, with a concomitant decrease in Kþ secretion. Unfortunately, this factor is still unidentified. 3.10.3.9. Serotonin and Other Biogenic Amines

The biogenic amine serotonin (5-hydroxytryptamine; 5HT) is known to stimulate diuresis in a variety of insects, including R. prolixus (Maddrell et al., 1969), L. migratoria (Morgan and Mordue, 1984; cAMP independent), P. brassicae (Nicolson and Millar, 1983), and M. sexta (Skaer et al., 2002). In most cases, levels required for stimulation are high: in R. prolixus, it stimulates distal tubules up to 1000-fold via a cAMP dependent mechanism with EC50
30–40 nmol l1 (Maddrell et al., 1993a), whereas the response of L. migratoria tubules is only 25% of the maximum obtained with a CC extract and is mediated by a different second messenger, possibly Ca2þ (EC50 10–100 nM; Morgan and Mordue, 1984). The response of A. aegypti tubules to serotonin varies with the stage of development. In larvae, it acts via cAMP to stimulate maximal secretion (EC50
100 nM; Clark and Bradley, 1996; Clark et al., 1998a). In contrast, the response of adult tubules is
25% of maximal (EC50
1 mM) and both cAMP and inositol trisphosphate (IP3) production are increased (Veenstra, 1988; Cady and Hagedorn, 1999b). In M. sexta four concentrations were studied; while no EC50 value was reported, it appears to be in the range of 100–300 mmol l1 (Skaer et al., 2002). Octopamine also stimulates M. sexta tubules, but is much less potent. Fournier et al. (1994) reported that serotonin has antidiuretic effects in the rectum of L. migratoria. Recently, Blumenthal (2003) has shown tyramine to be a potent stimulant of D. melanogaster MT, acting at levels as low as 1 nmol l1. Tyramine is several orders of magnitude more active than octopamine or dopamine. Interestingly, tyramine is produced in the MT from tyrosine taken up from the hemolymph, or culture medium, via tyrosine decarboxylase, and a tyramine receptor is also expressed in MT of this species. Tyramine is believed to play an autocrine or paracrine role; it causes fluctuations in the basolateral membrane transepithelial potential.

518 Hormones Controlling Homeostasis in Insects

Clearly this discovery merits studies in other species; this is the first demonstration of a physiological role for a tyramine receptor in D. melanogaster (Blumenthal, 2003). 3.10.3.10. Antidiuretic Factors Inhibiting Malpighian Tubule Secretion

Spring et al. (1988) characterized an antidiuretic hormone from hemolymph of A. domesticus kept under very dry conditions; an extract of this hemolymph inhibited secretion by MT of control insects. Hemolymph extract from insects reared under normal humidity did not inhibit MT secretion. Until recently, no factor had been isolated with such activity. Lavigne et al. (2001) published a partial purification of a factor from Leptinotarsa decemlineata through five RPLC steps using only two different columns. The protocol they used involved minimal changes of parameters between successive purifications, and utilized evaporation steps between RPLC purifications likely to result in poor recovery. While no peak was visible in the final separation step, they reported that the apparent molecular size of the factor (estimated by dialysis) was in the range of 25–50 amino acids. The first factor to be identified that inhibits MT secretion, Tenmo-ADFa, was isolated from pupal heads of T. molitor, based on its ability to elevate cyclic GMP (cGMP) levels in MT of this species. Tenmo-ADFa was found to be a 14-mer with the sequence VVNTPGHAVSYHVYOH, lacking the amidation usually found in regulatory peptides. It is exquisitely potent with an EC50 of 10 fmol l1 (0.01 pmol l1) (Eigenheer et al., 2002), but receptor downregulation occurs at high concentrations (such as 1 nmol l1!). A second factor isolated from the same source, Tenmo-ADFb, is a 13-mer with the sequence YDDGSYKPHIYGFOH, again nonamidated. It is 24 000 times less potent than Tenmo-ADFa, but still has EC50 ¼ 0.24 nmol l1 (Eigenheer et al., 2003). These factors have poor solubility in the acidic solvents usually used for extracting neuropeptides from tissues.

Following typical isolation protocols resulted in low yields. However, upon discovering that the bulk of the activity remained in the pellet from acid extraction, the pellet was extracted with a neutral solvent, which by then had so many impurities removed that only three RPLC purification steps were required to purify the factors from the neutral extract to homogeneity. Curiously, these factors have extremely high sequence similarity to two cuticular proteins of this beetle: Tenmo-ADFa is identical to the C-terminus of the protein CAA03880 (Mathelin et al., 1998) at all residues, except that Thr4 in the peptide is an Ala residue in the protein (Figure 9). Tenmo-ADFb is 100% identical with the 13 C-terminal residues of T. molitor putative cuticle protein 9.2 (TmPCP9.2) (Baernholdt and Andersen, 1998). Consequently, the identification of ADFb was not published until immunohistochemical evidence showed specific staining for Tenmo-ADFb-like material in two pairs of bilaterally symmetrical neurosecretory cells of the protocerebrum of T. molitor (Eigenheer et al., 2003). While the cuticle protein TmPCP9.2 lacks obvious enzymatic cleavage sites upstream of the portion identical with ADFb, this identity suggested the possibility that it could be a proteolysis fragment of the cuticle protein, and that this fragment might have coincidental biological activity. Tenmo-ADFa does have an interesting sequence identity to big endothelin 1 (Figure 9). The sequence identity to rabbit endothelin 1 (8 residues) and human endothelin 1 (7 residues) begins at exactly that bond where endothelin converting enzyme cleaves big endothelin into the potent vasoconstrictor endothelin 1. This cleavage site is marked with an arrow in Figure 9. Recently Coast (unpublished data) has found that M. sexta allatotropin (Kataoka et al., 1989a; Manse-AT) inhibits the Manse-DH stimulated secretion of fluid by larval MT of this species at 50–200 nmol l1 levels. This is the same concentration found earlier to inhibit fluid uptake from the anterior midgut (Lee et al., 1998) of M. sexta.

Figure 9 Two antidiuretic factors from Tenebrio molitor: Tenmo-ADFa and Tenmo-ADFb. Tenmo-ADFa is identical at all but one residue with the C-terminus of the cuticle protein CAA03880 from this species (Mathelin et al., 1998), and Tenmo-ADFb is 100% identical with the 13 C-terminal residues of another cuticle protein from this species, TmPCP9.2 (Baernholdt and Andersen, 1998). In addition, Tenmo-ADFa has an interesting sequence identity with big endothelin 1, a precursor of the potent vasoconstrictor endothelin 1. Endothelin converting enzyme cleaves big endothelin at the bond indicated by an arrow in the figure; this is exactly where the sequence identity with ADFa commences. Thus, the identity is to the apparently inactive endothelin fragment removed in processing.

Hormones Controlling Homeostasis in Insects

Incubation of 1 nmol l1 Manse-DH with and without 50 nmol l1 Manse-AT (E. Chidembo and D.A. Schooley, unpublished data) shows that Manse-AT seems to act via lowering cAMP levels. 3.10.3.11. Antidiuretic Factors that Promote Fluid Reabsorption in the Hindgut

Hormonal control of fluid reabsorption in the hindgut has been heavily studied in locusts (S. gregaria and L. migratoria). Three neuropeptides have been either isolated or characterized from the CC of locusts (reviews: Phillips et al., 1988, 1986): neuroparsins, ion transport peptide (ITP), and chloride transport stimulating hormone (CTSH). Only the first two have been fully sequenced; CTSH is labile to the usual conditions used for RPLC isolation of peptides, due to its instability in acidic conditions. Herault et al. (1985) reported that the nervous and glandular lobes of the CC of L. migratoria each contain a factor acting as an ADH on the rectum; these factors differ in size and extraction properties. The glandular lobe (GCC) factor was not purified, but Herault and Proux (1987) reported that GCC extracts cause a peak in rectal tissue cAMP levels coinciding with elevated short circuit current (Isc). This stimulation is mimicked by forskolin, a stimulant of adenylate cyclase. The factor from the CC was reported to be a neuroparsin (Np), because all antidiuretic activity in crude CC of L. migratoria was abolished by an antibody to this neuropeptide (Herault et al., 1988). Neuroparsins are proteins (NpA, NpB) isolated and sequenced from CC of L. migratoria (Girardie et al., 1989, 1990). They have also been reported to have hypertrehalosemic (Moreau et al., 1988) and antijuvenile hormone activity (Girardie et al., 1987). NpB was reported to be a homodimer of a 78-residue polypeptide. NpA is identical to NpB except for having a heterogeneous N-terminus, the longest form of which has 83 residues. NpB is thought to be formed from NpA by enzymatic cleavage of the N-terminal sequence. Subsequently Hietter et al. (1991) confirmed the sequences of these small proteins, including the N-terminal heterogeneity, but revised the structural assignment for neuroparsins involving three internal disulfide bridges, in which the chains are monomeric. Fournier (1991) presented a body of data consistent with NpB acting on rectal fluid reabsorption by stimulating the inositol phosphate (IP) cascade with subsequent elevation of cytosolic Ca2þ. They concluded that neuroparsins are the only ADH in L. migratoria GCC and that these peptides act via the IP-Ca2þ second messenger system, whereas serotonin from the NCC acts on the rectum via a cAMP mediated Ca2þ increase

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(Fournier et al., 1994). They did not study actions of purified or synthetic Nps on rectal solute transport processes. Jeffs and Phillips (1996) found no effect of Locmi-Nps on either rectal ion or fluid transport in the locust S. gregaria. They attributed this either to fundamental differences in transport properties in these closely related insects or, more likely, to the use of Cl-free saline in the assay used (Fournier et al., 1987) for preincubation of everted rectal sacs for 1 h. Chloride transport stimulating hormone, a peptide occurring in the CC of S. gregaria, was characterized by its ability to increase short circuit current in S. gregaria rectal sheets maintained in Ussing chambers (Phillips et al., 1980). In this assay, either cAMP or CC extracts give a ten-fold increase in Isc for >8 h after stimulation. A single zone is seen on size exclusion chromatography with Mr of about 8000 Da. The peptide nature is proved by its sensitivity to trypsin, and it is active at what appear to be nmol l1 concentrations (Phillips et al., 1986). The peptide can be extracted with water, saline, and aqueous ethanol, but biological activity is lost rapidly below pH 6. This may be consistent with the presence of an Asp-Pro bond in the primary sequence, which, as mentioned in Section 3.10.3.3, is suspected of causing an inability to isolate two CRF-like DH by RPLC techniques as well. Proux et al. (1985) provided some evidence that CTSH is produced in the pars intercerebralis and is transferred down nerves to the glandular lobe of the CC, after passing through the nervous lobe of the CC. CTSH is similar in size to Locmi-Nps, but is distinguished from the latter by the stability of Locmi-Nps to acidic conditions. Also, Locmi-Nps are reported to act via stimulation of phospholipase C. Audsley and Phillips (1990) used an assay similar to that for CTSH, substituting locust ileum for rectum, and found stimulants of ion transport in both the CC and the ventral ganglia. The effects elicited by CC and ganglia extracts differ in properties. The factor in the CC, unlike CTSH, is stable to acid. This stability led to its successful isolation by RPLC, and a partial sequence was obtained to 31 amino acids (Audsley et al., 1992b). This peptide was christened ion transport peptide (Schgr-ITP), to distinguish it from CTSH. It was found to be 44–59% identical to the crustacean hyperglycemic (CHH), molt inhibiting (MIH), and vitellogenesis inhibiting (VIH) hormones, a group of highly related 72 amino acid peptides (Audsley et al., 1992b). This was the first direct evidence for the existence of a peptide related to this family outside crustaceans. Audsley et al. (1992a) tested purified Schgr-ITP on the rectum and found that it had weak effects on short circuit

520 Hormones Controlling Homeostasis in Insects

current compared with CTSH, and no effect on the rate of fluid reabsorption. They concluded that separate neuropeptides act on the ileum and rectum of the locust. Meredith et al. (1996) used degenerate primers designed from the partial protein sequence to clone by PCR a partial sequence of Schgr-ITP from a brain cDNA library. Use of 50 and 30 rapid amplification of cDNA ends (RACE) strategies led to isolation of a cDNA of 517 bp encoding a complete open reading frame for Schgr-ITP prepropeptide of 130 amino acid residues (Meredith et al., 1996). The prepropeptide has a 55-residue signal peptide and a dibasic cleavage site preceding the start of the partial ITP sequence determined by Audsley et al. (1992b). The C-terminal Leu is followed by Gly-Lys-Lys-stop, which is consistent with C-terminal amidation and a second dibasic cleavage site to give the complete ITP sequence of 72 amino acid residues. Meredith et al. (1996) also used ITP cDNA sequence primers to probe a locust ileal mRNA library, and isolated an ITP-like clone, which was sequenced. It was identical to the brain cDNA for ITP except for an additional 121 bp insert at amino acid position 40 of ITP, suggesting alternative C-termini splicing of genomic DNA. The open reading frame for ITP-L (134 residues) is four residues longer than that of ITP (Figure 10). All six cysteines are conserved. The unique C-terminus of ITP-L has only 14 of the last 36 amino acid residues in common with ITP, most of the difference being over the last 20 residues. Using reverse transcriptase PCR (RT-PCR) techniques, ITP-L mRNA was detected in tissues (flight muscle, hindgut, and MTs) whose extracts have no stimulatory effect in the locust ileal Isc bioassay, while ITP mRNA was found only in the brain and CC, which do stimulate ileal Isc. The ITP-L peptide does not stimulate the locust ileal Isc bioassay, but acts as a weak antagonist (Wang et al., 2000). The Schgr-ITP synthesized by solid-phase peptide synthesis was

oxidized to the appropriately folded form with three disulfide bonds (King et al., 1999); this was found to exhibit biological actions identical to that of the native peptide isolated from tissue. While in the 1990s there was concern over conflicts in results between the proponents of neuroparsins versus Schgr-ITP, three publications in the area of crustacean neuroendocrinology strongly support the role of ITP as an authentic osmoregulator (see Chapter 3.16). Chung et al. (1999) reported a large, precisely timed release of CHH from gut endocrine cells in the crab Carcinus maenas at ecdysis. This release appears to trigger the water and ion uptake required during molting, which allows the swelling necessary for successful ecdysis and the subsequent increase in size during postmolt. The endogenous ortholog of CHH of the freshwater crayfish Pachygrapsus marmoratus (SpaningsPierrot et al., 2000) has been shown to play a crucial role in control of gill ion transport. In the crayfish Astacus leptodactylus, injection of CHH was found to increase the hemolymph osmolality and Naþ concentration 24 h after injection (Serrano et al., 2003). Two other CHH related peptides caused a smaller increase in Naþ concentration. Liao et al. (2000) characterized one of two factors that separate on RPLC of brain–CC–corpora allata extracts of larval M. sexta. These factors trigger fluid reabsorption in an everted rectal sac bioassay with M. sexta. Owing to the cryptonephric anatomy of larvae of this species, the CRF-like peptide ManseDH also causes rectal fluid reabsorption. However, the effect of Manse-DH is blocked by bumetanide (an inhibitor of the Naþ–Kþ–2Cl cotransporter), bafilomycin A1, and amiloride, while that of the more potent of the two factors, Manse-ADFB, is not blocked by any of these inhibitors. The Cl channel blockers 4,40 -diisothiocyanatostilben e-2, 20 -disulforic acid disodium salt (DIDS) and diphenylamine-2-carboxylic acid (DPC) both block

Figure 10 Alignment of Schgr-ITP with its Bombyx mori (Bommo-) and Drosophila melanogaster paralogs (Clustal W, V. 1.8), as well as the variant form Schgr-ITPL. Boxed residues are identical. Also shown are the highly similar Carcinus maenas (Carma-) crustacean hyperglycemic hormone, and Homarus americanus (Homam-) molt inhibiting hormone. All peptides are amidated at the C-terminus (denoted by lower case a); the two crustacean peptides are blocked at the N-terminus with pQ (pyroglutamate) residues.

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the action of Manse-ADFB, as does H-89, a potent and specific inhibitor of protein kinase A. These data suggest that this factor of still unknown structure has a mechanism of action similar to Schgr-ITP. Homologs of ITP have been identified in B. mori (Endo et al., 2000) and in the genome of D. melanogaster (Figure 10): While neither has been synthesized or subjected to an activity assay, combined with the data on osmoregulatory effects of CHH, it seems likely that Manse-ADFB is similar to this family of peptides. 3.10.3.12. Cellular Mechanisms of Action

3.10.3.12.1. Introduction Does the diverse control of tubule secretion between different orders, and even between similar species belonging to the same order, have any rationale? Do differences in hemolymph composition, diet (and hence ion uptake), water availability, and the structure of the excretory system explain the diverse control strategies employed by different species? Primitive insect orders have a hemolymph characterized by a high concentration of Naþ relative to Kþ, and the sum of total anions and cations accounts for a considerable part of the total osmolarity (Sutcliffe, 1963). In contrast, more highly evolved insect orders have a greater proportion of hemolymph osmolarity attributable to high levels of amino acids and organic acids and, in some (Lepidoptera, Hymenoptera, and certain Coleoptera; Sutcliffe, 1963), Kþ is the dominant cation, with elevated concentrations of Mg2þ and Ca2þ. Since tubule secretion is driven by secondary active cation transport, differences in hemolymph composition might place some constraint on how fluid secretion is regulated. The same could apply to the dietary intake of ions, most notably Kþ and Naþ. If the diet is Kþ-rich, as in herbivorous insects, then it is appropriate for Kþ to be the dominant cation secreted by the MT and to control secretion by manipulating the rate of Kþ transport. Conversely, hematophagous insects take infrequent blood meals, which imposes a considerable challenge on the excretory system for the removal of imbibed salt (NaCl) and water. Not surprisingly, the MTs of mosquitos (A. aegypti) respond to released diuretics with a dramatic increase in secretion driven by a switch from Kþ transport to Naþ transport (Beyenbach, 2003). Hence, the diuretic hormone (so far unidentified, but thought to be a CRF-like or CT-like DH) exerts both diuretic and natriuretic activity. Interestingly, in the laboratory, the same mechanism has been shown to operate in the tubules of male mosquitos (Plawner et al., 1991), which do not feed on blood but on nectar (Clements, 1992), resulting in a diet

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that is not rich in NaCl. Periods of diuresis that do not require a concomitant natriuresis must be initiated by the release of a different DH, possibly a myokinin, which, because it stimulates anion movement, has a neutral effect on cation transport. Can a similar story be developed for other species? D. melanogaster feeds on fermentation products (the alcoholic of the Insecta!), whereas the closely related M. domestica imbibes a mixture of saliva and the products of extracorporal digestion. A major drawback in this analysis is that most researchers have studied effects on fluid secretion rather than on ion transport. Peptides with apparently similar diuretic activity could well differ in their effect on urine composition and hence on ion transport (the product of concentration and the rate of secretion). An early demonstration of the possibility of such an occurrence is in MT of A. aegypti treated with bumetanide, a drug that inhibits the Naþ–Kþ–2Cl cotransporter stimulated by cAMP. Bumetanide treatment has little effect on basal fluid secretion, but decreases Kþ secretion with a concomitant increase in Naþ secretion (Hegarty et al., 1991). Coast (1995) has shown that Locmi-DH (CRF-like) and locustakinin have fairly similar diuretic potency, but have quite different effects on ion transport. The majority of physiological studies to date on the action of synthetic peptides affecting MT function have focused on cellular effects, such as elevation of second messengers, or on fluid transport. Relatively few have focused on the effects of synthetic factors on ion transport, which, given the complex interplay of factors controlling fluid and ion homeostasis in vertebrates, is a serious omission. Moreover, in the cricket Teleogryllus oceanicus, the distal portion of the MT have been reported to secrete a hyperosmotic urine containing 125 mM Mg2þ, together with a lesser concentration of Naþ (Xu and Marshall, 1999). There are few studies on Mg2þ transporters in insect systems, yet in a number of insect species with high hemolymph Kþ, the concentration of Mg2þ is actually higher (Sutcliffe, 1963; Cohen and Patana, 1982; Dow et al., 1984). Thus, data on specific ion transport is an underinvestigated area likely to shed some light on why so many peptides are involved in control of water and salt balance in insects. Water availability may also play a role in determining the cocktail of diuretics and antidiuretics used to control tubule secretion. The simplistic view that diuretics stimulate tubule secretion whereas antidiuretics increase fluid reabsorption in the hindgut is no longer tenable (at least in some species). To date, antidiuretic activity inhibiting the MT has been identified in three species. In R. prolixus,

522 Hormones Controlling Homeostasis in Insects

the antidiuretic activity of CAP2b has been linked to the need to rapidly turn off tubule secretion (Quinlan et al., 1997). Normally, the rapid degradation of a DH (probable half-life in the circulation measured in minutes) (Li et al., 1997) would suffice to end a period of diuresis. The situation in R. prolixus is very different, however, because the DH has such a dramatic effect on secretion, which at maximal rates would be sufficient to deplete the total store of water in the hemolymph within minutes. Under these conditions, it is essential that diuresis be switched off coincident with reduced fluid uptake from the midgut (Quinlan et al., 1997), and the insect now conserves water until the next blood meal. In this respect, R. prolixus resembles a xeric insect in the period between its intermittent blood meals. It is of considerable interest that another insect shown to use antidiuretic factors to control tubule secretion is T. molitor, an extreme example of a xeric species able to survive very dry conditions (O’Donnell and Machin, 1991). In both T. molitor and R. prolixus (for which a native antidiuretic factor has still to be identified), antidiuresis appears to result from the activation of a cGMP dependent cAMP phosphodiesterase, which lowers intracellular levels of cAMP, the second messenger used by both CRF-like and CT-like DH. Not surprisingly, the antidiuretic factors of T. molitor are extremely potent (Eigenheer et al., 2002, 2003) and effectively counter the diuretic activity of native CRF-like DH (Wiehart et al., 2002). To grasp the importance of this, it is important to note that coleopteran larvae and adults have a cryptonephric system in which the distal ends of the MTs are closely associated with the rectum (Ramsay, 1964; Section 3.10.3.12.7). Fluid is reabsorbed from the rectum into the cryptonephric region of the tubules and flows into the gut at the midgut–hindgut junction, where it is postulated to move anteriorly directed and to moisten the dry food for digestion prior to reabsorption by the midgut (Nicolson, 1991). To gain water (the cryptonephric system is used to take up water from subsaturated atmospheres), fluid absorption from the free portion of the tubules must exceed fluid secretion, although these two processes could be taking place in functionally different segments. Unfortunately, nothing is known of fluid reabsorption from T. molitor tubules or whether antidiuretic factors have any effect on urine composition. Recent work (G.M. Coast, unpublished data) with MT from M. sexta larvae, which also have a cryptonephric system, shows that one segment (the yellow segment) can display net secretion or net reabsorption depending on the composition of the bathing fluid and the complement of peptides present. The two

processes appear to be proceeding in parallel, with the overall balance tilted towards secretion or reabsorption by stimulants of each activity. The likelihood must be that similar mechanisms operate in T. molitor tubules. 3.10.3.12.2. CRF-like DH:cellular actions The CRF-like DHs studied to date all elevate cAMP in response to treatment of MT in vitro. As noted by Rafaeli et al. (1984), the cAMP is usually released from the tubule, which allows assay of the medium without having to homogenize and extract the tubules. This fact facilitates in vitro assays for isolating new members of this family. When using conspecific assays, CRF-like DH stimulate secretion by MTs of A. domesticus (Coast and Kay, 1994), L. migratoria (Patel et al., 1995), D. punctata (Furuya et al., 2000b), M. domestica (Iaboni et al., 1998), T. molitor (both Tenmo-DH37 and TenmoDH47) (Wiehart et al., 2002), and both CRF-like DH of M. sexta, Manse-DH (Audsley et al., 1993, 1995), and Manse-DPII (Blackburn and Ma, 1994) at low nanomolar concentrations. Diuretic activity varies among species, from a maximal response in A. domesticus (Coast and Kay, 1994), equivalent to that obtained with a CC extract, to 60% Z11–14:CoA indicate that these enzymes have the inherent specificity to produce the 92/8 ratio of the major pheromone components Z11– and E11–14:OAc. Two minor pheromone components are produced by chain shortening Z11– and E11–14:OAc. The ratio of Z9– to E9–12:OAc is about 1 to 2. This indicates that the chain-shortening enzymes may prefer E11–14:CoA or that very little Z11–14:CoA is available to chain shorten. This combined information indicates that in A. velutinana pheromone glands, the final ratio of pheromone components can be produced through the concerted action of a D11-desaturase that produces at least a 60/40 ratio of Z/E intermediate isomers. The final ratio of acetate esters (92/8) is produced through the specificity for the Z isomer by the acetyltransferase. The minor components are produced by specificity in chain shortening. Another insect that utilizes specific ratios of Z11– and E11–14:OAc is the European corn borer, O. nubilalis. Two strains are known in which one produces a ratio of Z/E of about 97/3 (Z strain) and the other produces an opposite ratio of Z/E of about 1/99 (E strain). Hybridization studies between the two strains indicated that offspring have an acetate

ester ratio of Z/E of about 30/70 (Klun and Maini, 1982). The D11-desaturase from both strains produced a product with about 30/70 Z/E in an in vitro enzyme assay (Wolf and Roelofs, 1987). These results indicate that the final ratio of acetate ester isomers is produced after the desaturation step. The enzymes that follow the desaturase are a reductase to make an alcohol and an acetyltransferase to produce the acetate esters. Two studies have shown that the acetyltransferase is similar between the two strains (Jurenka and Roelofs, 1989; Zhu et al., 1996). However, labeled acids applied to glands in vivo were selectively incorporated into the correct pheromone ratio indicating that the reductase shows specificity (Zhu et al., 1996). Therefore, the final pheromone ratios produced by females of the European corn borer are made through the action of a D11-desaturase that can produce both Z and E isomers. The final acetate ester ratio is strain dependent and is produced through the specificity found in the reductase system. The above three examples illustrate how a species-specific pheromone blend is produced by the concerted action of desaturases, chain-shortening enzymes, and a reductase and an acetyltransferase. The specificity inherent in certain enzymes in the pathway produces the final blend of pheromone components. 3.14.4.3.6. Hydrocarbon pheromones Moths in the families Geometridae, Arctiidae, Amatidae, Lymantriidae, Lyonetiidae, and some Noctuidae utilize hydrocarbons or epoxides of hydrocarbons as their sex pheromones. Biosynthesis of hydrocarbons occurs in oenocyte cells that are associated with either epidermal cells or fat body cells (Romer, 1991). Once the hydrocarbons are biosynthesized they are transported to the sex pheromone gland by lipophorin (Schal et al., 1998a). When the transport of hydrocarbon sex pheromones in arctiid moths was investigated in detail by Schal et al. (1998a), it was found that a very specific uptake was occurring at pheromone glands. Lipophorin was shown to contain both the sex pheromone and cuticular hydrocarbons; however, only the pheromone gland had the sex pheromone. Other studies have shown similar pathways in other moths (Jurenka and Subchev, 2000; Subchev and Jurenka, 2001; Wei et al., 2003). Most moth sex pheromones that are straight chain hydrocarbons also usually have an odd number of carbons. Most of these are polyunsaturated with double bonds in the 3,6,9- or 6,9-positions, indicating that they are derived from linolenic or linoleic acid, respectively (Rule and Roelofs, 1989;

Biochemistry and Molecular Biology of Pheromone Production

Millar, 2000). Linolenic and linoleic acid cannot be biosynthesized by moths so they must be obtained from the diet (Stanley-Samuelson et al., 1988). A few even-chain-length hydrocarbon sex pheromones have been identified that also have 3,6,9- or 6,9-double bond configurations (Millar, 2000), indicating that they too are derived from linolenic or linoleic acids; however, it is not known how these even-chain hydrocarbons are formed. A major class of sex pheromones that are derived from hydrocarbons are the polyene monoepoxides (Millar, 2000). These usually have double bonds in the 3,6,9-positions or 6,9-positions, again indicating that they are biosynthesized from linolenic or linoleic acids, respectively. Although the production of hydrocarbon occurs in oenocytes the epoxidation step takes place in the pheromone gland. This has been demonstrated in several studies utilizing deuterium labeled precursors. In a study on the Japanese giant looper, Ascotis selenaria cretacea, that uses 6,9–19:3,4Epox as a sex pheromone component, deuterium-labeled hydrocarbon precursor, D3–3,6,9–19:Hc, was topically applied to pheromone glands and found to be converted to the epoxide, indicating that epoxidation takes place in pheromone glands (Miyamoto et al., 1999). By using a variety of polyene precursors, it was also determined that the monooxygenase regiospecifically attacked the n-3 double bond regardless of chain length or degree of unsaturation. This indicates that the epoxidation enzyme is regiospecific in this insect (Miyamoto et al., 1999). A study using the gypsy moth, L. dispar, illustrates the overall pathways involved in production of

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epoxide pheromone components (Figure 6) (Jurenka et al., 2003). This insect uses disparlure, 2me18:7,8Epox, as a pheromone component. Incubation of isolated abdominal epidermal tissue with deuterium-labeled valine resulted in incorporation into 2me-Z7–18:Hc. This indicates that the oenocyte cells associated with the epidermal tissues biosynthesize 2me-Z7–18:Hc using the carbons of valine to initiate the chain. The double bond is probably introduced by a D12-desaturase as determined by using specific deuterium-labeled intermediates. Hemolymph transport of 2me-Z7–18:Hc is indicated by the finding of this alkene in the hemolymph (Jurenka and Subchev, 2000). Demonstration that 2me-Z7–18:Hc is converted to the epoxide in the pheromone gland was shown by using deuterium-labeled 2me-Z7–18:Hc and incubation with isolated pheromone glands. Disparlure is a stereoisomer that has the 7R,8S or (þ) configuration and chiral chromatography indicated that only the (þ)-isomer was produced by pheromone glands (Jurenka et al., 2003). These results indicate that hydrocarbon pheromones and their epoxides are produced through a pathway outlined in Figure 6. 3.14.4.4. Pheromone Biosynthesis in Beetles

Pheromone biosynthesis in the Coleoptera is as diverse as the taxa and the pheromone structures, and the utilization of several types of pheromone biosynthetic pathways has been demonstrated (Vanderwel and Oehlschlager, 1987; Vanderwel, 1994; Seybold and Vanderwel, 2003; Tittiger, 2003). Extensive work has been done on the biosynthesis of coleopteran pheromones, and the major

Figure 6 Production of the sex pheromone in the gypsy moth, Lymantria dispar. The oenocyte cells located in the abdomen biosynthesize the alkene hydrocarbon precursor to the pheromone, 2me-Z 7–18:Hc. It is transported through the hemolymph by lipophorin. The alkene is taken up by pheromone gland cells where it is acted upon by an epoxidase to produce the pheromone disparlure, 2me-18:7,8Epox. (Reprinted with permission from Jurenka, R.A., 2003. Biochemistry of female moth sex pheromones. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Pheromone Biochemistry and Molecular Biology. Elsevier, San Diego, CA, pp. 53–80; ß Elsevier.)

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systems that have been investigated are described below. Beetles can generate pheromone either by modification of dietary host compounds or de novo biosynthesis, with the latter accounting for the majority of beetle pheromone components.

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3.14.4.4.1. Isoprenoid pheromones from bark beetles Most of the knowledge about beetle pheromone biosynthesis and endocrine regulation (see Section 3.14.4.5.2) comes from studies of various bark beetles, especially Ips and Dendroctonus species (Scolytidae). Some bark beetles may modify fatty acyl or amino acid precursors (Vanderwel and Oehlschlager, 1987; Birgersson et al., 1990); however, the majority of pheromone components are isoprenoid (Schlyter and Birgarson, 1999: Seybold et al., 2000). Understanding of the origin of bark beetle pheromone components has undergone a paradigm shift in the last decade. Until recently, it was widely accepted that bark beetles, in contrast to most other insects (Tillman et al., 1999), obtained their pheromone components by simple modification of of host tree dietary precursors (reviews: Bordon, 1985; Vanderwel and Oelschlager, 1987; Vanderwel, 1994). This model was based on various studies that showed the association of monoterpene synthesis with conifers (Croteau, 1981), the abundance of potential precursors in the host tree (Gershenzon and Croteau, 1991), the demonstration that monoterpenoid precursors such as myrcene and a-pinene could be converted to the pheromone components ipsdienol, ipsenol, and cis and trans verbenol (Hughes, 1974; Renwick et al., 1976a, 1976b; Byers, 1981), and the conclusive demonstration that deuterated myrcene was converted to ipsenol and ipsdienol in I. paraconfusus (Hendry et al., 1980). In the past decade and a half, however, significant evidence has emerged supporting the de novo biosynthesis of most bark beetle pheromone components. First, doubts were raised about the role of myrcene in ipsdienol and ipsenol production when it was noted that myrcene may not be present in sufficient quantity in some host trees to account for the amount of pheromone that was produced (Byers, 1981; Byers and Birgersson, 1990). Second, Ips beetles treated with the HMG-R reductase inhibitor compactin show a marked decrease in ipsdienol production (Ivarsson et al., 1993). Third, myrcene-treated male I. pini produce ipsdienol with a racemic enantiomeric composition whereas JH III-treated males produce ipsdienol with a 87– 96% () enantiomeric composition (Lu, Blomquist, and Seybold, unpublished data). The naturally occurring enantiomeric composition of ipsdienol

from California I. pini is 95–98% () (Seybold et al., 1995a). Fourth, JH III-treated male I. pini incorporate labeled acetate and mevalonate into ipsdienol (Seybold et al., 1995b; Tillman et al., 1998). 2-Methyl-3-buten-2-ol is similarly synthesized de novo in I. typographus (Lanne et al., 1989). More recently, key genes in pheromone production, including HMG-R and HMG-CoA synthase (HMG-S) have expression patterns consistent with their roles in regulating de novo isoprenoid pheromone biosynthesis (Tittiger et al., 1999; Tillman, Blomquist, and Seybold, unpublished data). A geranyl diphosphate synthase (GPPS) cDNA from I. pini was also isolated, functionally expressed, and modeled (Gilg-Young, Welch, Tittiger, and Blomquist, unpublished results). The existence of this novel enzyme argues strongly for the evolution of de novo pheromone biosynthetic capacity in bark beetles. Taken together, the data emerging from Ips species overwhelmingly indicate the de novo production of monoterpenoid pheromone components. These data are supported by studies in other beetles that similarly prove or imply de novo pheromone component biosynthesis. Radiotracer studies demonstrated the de novo biosynthesis of frontalin in a number of Dendroctonus species (Barkawi et al., 2003). Expression patterns of HMG-R and HMG-S in D. jeffreyi correlate tightly with frontalin production (Tittiger et al., 2000, 2003). The four monterpene alcohols and aldehydes in the boll weevil, A. grandis, are also synthesized from mevalonate and acetate (Mitlin and Hedin, 1974; Thompson and Mitlin, 1979). The capacity for de novo biosynthesis does not preclude the conversion of host precursors to pheromone components. For example, the incorporation of acetate and mevalonolactone into ipsdienol and ipsenol may proceed through the conversion of geranyl diphosphate to myrcene, which could be directly hydroxylated to ipsdienol and E-myrcenol, and indirectly, perhaps through a ketone intermediate, to ipsenol (Martin et al., 2003). In this scheme, host myrcene ingested during feeding would enter the de novo pathway downstream of geranyl diphosphate. Similarly, cotton plant monoterpenes (myrcene and limonene) could enter a de novo biosynthetic pathway to grandlure in A. grandis. The question then arises as to whether de novo biosynthesis or host precursor conversion is the preferred route to pheromone production. Male I. pini exposed to myrcene vapors produce a racemic mixture of ipsdienol, whereas the naturally occurring pheromone of Western I. pini is about 95 : 5 ()/(þ) ratio (Lu, Blomquist, and Seybold, unpublished data). This suggests that myrcene is not the

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direct precursor to ipsdienol, but that its hydroxylation is a true detoxification reaction. Perhaps a highly specific hydroxylase may synthesize ipsdienol de novo, while a hydroxylase with a different specificity detoxifies myrcene. Other possibilities also exist, particularly considering that ipsdienone appears to be a precursor to ipsdienol (Ivarsson et al., 1997). To make ipsdienol, it is possible that geranyl diphosphate is hydroxylated and then dephosphorylated, bypassing myrcene as an intermediate. Thus, though the final biosynthetic steps remain uncharacterized, de novo biosynthesis is

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clearly the most important route to ipsdienol and ipsenol in I. pini and I. paraconfusus. Further examples should arise as other species are studied. A proposed biosynthetic scheme for a number of coleopteran isoprenoid pheromone components is presented in Figure 7. Hemiterpene pheromone components of the bark beetles are similarly synthesized de novo. Lanne et al. (1989) demonstrated the incorporation of labeled acetate, glucose, and mevalonate into 2-methyl-3-buten-2-ol in I. typographus. This also argues for the de novo synthesis of 3-methyl-3-buten-1-ol and 3-methyl-2-buten-1-ol.

Figure 7 Proposed biosynthetic pathways for a number of hemi- and monoterpenoid pheromone components in the Coleoptera.

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The mevalonate pathway intermediate, dimethylallyl diphosphate, likely provides the carbon skeleton for 3-methyl-2-buten-1-ol by dephosphorylation. The other five-carbon intermediate, isopentenyl diphosphate, could be directly converted to 3-methyl-3-buten-1-ol, and perhaps through several steps, to 2-methyl-3-buten-2-ol. Ipsdienol and ipsenol production from geranyl diphosphate appears to involve relatively straightforward though not yet fully characterized biochemical modifications. The production of frontalin, and the recently identified male-produced pheromone of the Colorado potato beetle, Leptinotarsa decemlineata (S)-3,7-dimethyl-2-oxo-oct-6-ene-1,3-diol (CPB1) (Dickens et al., 2002), is less clear. Barkawi et al. (2003) demonstrated that acetate and mevalonolactone are precursors to frontalin, proving the isoprenoid origin of this common Dendroctonus pheromone component. he carbon skeleton of frontalin probably arises from geranyl disphosphate via a putative dioxygenase, which converts geranyl diphosphate to sulcatone. Sulcatone is an obvious precursor to sulcatol, which is a pheromone component of some bark beetles (Gnathotrichus sulcatus) (Byrne et al., 1974). In Dendroctonus spp., geranyl diphosphate-derived sulcatone may be converted to 6-methyl-6-hepten-2-one (6-MHO), a known intermediate to frontalin (Perez et al., 1996). Alternative cyclizations of 6-MHO in other beetles could lead to the pheromone components pityol and vittatol. Significant amounts of sulcatone are also found in Colorado potato beetle males during their synthesis of CPB1 (Dickens et al., 2002). It is easy to envision CPB1 as an intermediate in the dioxygenase reaction (Figure 7). The picture emerging from these studies is that isoprenoid pheromone component production in beetles is mostly de novo, with carbon being diverted from the mevalonate pathway at geranyl diphosphate. Geranyl diphosphate may be directly modified through dephosphorylation and cyclizations (cotton boll weevil) or hydroxylations (Ips spp.). Alternatively, some bark beetles, such as Dendroctonus spp., apparently have a dioxygenase that oxidizes geranyl diphosphate or geraniol to produce a ketodiol (CPB1) or sulcatone, which acts as a precursor to some pheromone components such as sulcatol, pityol, and frontalin. 3.14.4.4.2. Fatty acid-derived pheromones Numerous beetle genera use modified fatty-acyl compounds as pheromone components. Less is known about their biosynthesis compared to that of isoprenoid pheromones, but the same general strategy

of modifying or combining existing biosynthetic pathways is conserved. For some beetles, the modifications are relatively minor. For example, Attagenus spp. (Dermestidae) myristic acid may be desaturated at the D5 and D7 positions to produce tetradecadienoic acid pheromone components. The stereochemistries of the double bonds apparently provide specificity between species (Fukui et al., 1977). It is unclear whether the short-chain fatty acid precursors to these pheromone components are synthesized through normal fatty acid elongations, or are the b-oxidation products of longer fatty acids. For other beetles, modifications can become more complex. Female Tenebrio molitor produce 4methyl-1-nonanol from propionyl-, malonyl-, and methylmalonyl-precursors (Islam et al., 1999). This is an example of carbon being shunted away from fatty-acyl elongation before long fatty acids are completed. The use of methylmalonate to produce methyl-branched hydrocarbons is well established in other insect systems (Blomquist, 2003; Schal et al., 2003), though it is unknown if beetles have a secondary fatty acyl synthase which, similar to that in houseflies, incorporates methylmalonyl-CoA precursors efficiently. The flexibility of the fatty acid biosynthetic pathway is extended in some nitidulid beetles (Carpophilus spp.), where males use propionate and butyrate (presumably as methylmalonate and ethylmalonate) to make methyl- and ethyl-branched triene and tetraene pheromone components (Figure 1), apparently also via the fatty acid biosynthetic pathway (Bartelt et al., 1992). The branched hydrocarbons generally have 10–12 carbon backbones with conjugated double bonds. In contrast to other systems, where pheromone component biosynthesis is highly specific, Carpophilus spp. males produce a mixture of related structures, some of which act as pheromones and some of which do not. Since di-substituted tetraenes are less abundant than mono- or unsubstituted tetraenes, it appears that nonacyl units placed in the growing hydrocarbon chains represent ‘‘mistakes’’ made by a synthesis machinery with a low stringency for substrate selection (Bartelt, 1999). Such nonspecific hydrocarbon biosynthesis may serve speciation, since changes in antennal receptivity may accommodate preexisting compounds (Bartelt, 1999). Interestingly, the desaturated nature of these hydrocarbons is not due to fatty acyl desaturases, but due to the inactivity of enoyl-ACP reductase during biosynthesis so that the enoyl-ACP intermediate formed during elongation is not reduced (Petroski et al., 1994). This

Biochemistry and Molecular Biology of Pheromone Production

suggests that carbon is shunted out of the fatty acid biosynthetic pathway when the chains are of the correct length, similar to the situation in T. molitor. Rather than modifying the normal biosynthetic pathway to produce pheromone components, some beetles modify normal products of the pathway. For example, lactone pheromone components of some scarab beetles are produced by the stereospecific alterations of long chain fatty acids. Female Anomala japonica (Scarabaeidae) are perhaps best studied among scarab beetles for the biosynthesis of japonilure and buibuilactone, which involves the successive D9 desaturation, hydroxylation, two rounds of b-oxidation to shorten the chain length, and cyclization of stearic and palmitic acids (Leal et al., 1999). Of all these, only the hydroxylation step appears to be stereospecific (Leal, 1998). This step is important because different enantiomers have different functions in different Anomala species (Leal et al., 1999). 3.14.4.4.3. Host precursor modifications While most bark beetle isoprenoid pheromones are clearly synthesized de novo, there is strong evidence that some pheromone components are indeed the result of modifying host precursor molecules. a-Pinene is produced naturally by pine trees, and can be hydroxylated to cis- and trans-verbenol (Figure 1) by Ips beetles (Renwick et al., 1976a). A further oxidation of verbenol yields verbenone in Dendroctonus ponderosae (Hunt and Bordon, 1989). Similarly, Jeffrey pine trees contain relatively low levels of monoterpenoids, but high levels of heptane. Female Dendroctonus jeffreyi that attack these trees produce 1-heptanol and 2-heptanol, and 1-heptanol acts as a sex pheromone (Paine et al., 1999). 3.14.4.5. Pheromone Biosynthesis in Diptera

3.14.4.5.1.1. Housefly pheromone biosynthesis: C23 sex pheromone components A combination of in vivo and in vitro studies using both radio- and stable-isotope techniques established the biosynthetic pathways for the major sex pheromone components in the housefly (Figure 8) (Dillwith et al., 1981, 1982; Dillwith and Blomquist, 1982; Blomquist et al., 1984b; Vaz et al., 1988). (Z)-9-tricosene is formed by the microsomal elongation of 18 : 1CoA to 24 : 1-CoA using malonyl-CoA and NADPH, and the elongated fatty acyl-CoA is then reduced to the aldehyde and converted to the hydrocarbon one carbon shorter (Figure 8) (Reed et al., 1994, 1995; Mpuru et al., 1996). A cytochrome P450 enzyme is involved in the metabolism of the alkene to the corresponding epoxide and ketone

721

(Ahmad et al., 1987). It appears that the same enzyme that catalyzes formation of an epoxide from the double bond between carbons 9 and 10 of the alkene of (Z)-9-tricosene also hydroxylates it at position n-10. The secondary alcohol thus formed is then converted to the unsaturated ketone (Guo et al., 1991). 3.14.4.5.1.2. Mechanism of hydrocarbon formation: decarboxylation? The mechanism of hydrocarbon formation has proven elusive. In an elegant set of experiments in the 1960s and early 1970s, Kolattukudy and coworkers demonstrated that fatty acyl-CoAs were elongated and then converted to hydrocarbon by the loss of the carboxyl group (reviews: Kolattukudy et al., 1976; Kolattukudy, 1980). In the 1980s and early 1990s, the hypothesis was put forward that very long chain fatty acylCoAs were reduced to aldehyde and then decarbonylated to hydrocarbon and carbon monoxide, and that this reaction did not require any cofactors or O2. Evidence for this decarbonylation mechanism was obtained from a plant (Cheesbrough and Kolattukudy, 1984), an alga (Dennis and Kolattukudy, 1991, 1992), a vertebrate (Cheesbrough and Kolattukudy, 1988), and an insect (Yoder et al., 1992). In the work on hydrocarbon formation in the housefly, it was found that the acyl-CoA is reduced to the aldehyde, with the conversion of the aldehyde to hydrocarbon requiring NADPH and molecular oxygen, and that the products were hydrocarbon and carbon dioxide (demonstrated by radio-gas– liquid chromatography) (Reed et al., 1994, 1995). Antibodies to housefly cytochrome P450 and to P450 reductase inhibited hydrocarbon formation in microsomes, as did exposure to CO, and the latter could be partially reversed by white light. GC-MS analyses of specifically deuterated substrates showed that the protons on positions 2,2 and 3,3 of the acyl-CoA were retained during conversion to hydrocarbon, and that the proton on position 1 of the aldehyde was transferred to the adjacent carbon and retained during hydrocarbon formation. Furthermore, several peroxides could substitute for O2 and NADPH and support hydrocarbon production. All this evidence strongly supports a cytochrome P450 involvement in hydrocarbon synthesis (Reed et al., 1994). The mechanism of hydrocarbon formation remains controversial, with evidence obtained favoring both a decarbonylation and a decarboxylation mechanism. The resolution of the problem awaits cloning, expressing, and assaying the enzymes involved.

722 Biochemistry and Molecular Biology of Pheromone Production

Figure 8 Biosynthetic pathways showing the putative steps at which ecdysteroids regulate the hydrocarbon and hydrocarbonderived pheromone components of the female housefly, Musca domestica. (Reprinted with permission from Blomquist, G.J., 2003. Biosynthesis and ecdysteroid regulation of housefly sex pheromone production. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Pheromone Biochemistry and Molecular Biology. Elsevier, New York, pp. 131–252; ß Elsevier.)

3.14.4.5.1.3. Biosynthesis of the methylalkane pheromone components Methylalkanes are formed by the substitution of methylmalonyl-CoA in place of malonyl-CoA at specific points during chain elongation. Carbon-13 nuclear magnetic resonance (NMR), mass spectrometry, and

radiochemical studies (Dwyer et al., 1981; Dillwith et al., 1982; Chase et al., 1990) demonstrated that the methylmalonyl-CoA was added during the initial steps of chain elongation in insects using what appears to be a novel microsomal fatty acid synthase (FAS) (Figure 8). A microsomal FAS was

Biochemistry and Molecular Biology of Pheromone Production

first suggested from studies on T. ni for the formation of methyl-branched very long chain alcohols (de Renobales et al., 1989). In this insect, high rates of methyl-branched very long chain alcohol synthesis were observed in the mid-pupal stages at times when soluble FAS activity was very low or undetectable. The FAS of most organisms is soluble (cytoplasmic). A microsomal FAS was also implicated in the biosynthesis of methyl-branched fatty acids and methyl-branched hydrocarbon precursors of the German cockroach contact sex pheromone (Juarez et al., 1992; Gu et al., 1993). A microsomal FAS present in the epidermal tissues of the housefly is a likely candidate responsible for methyl-branched fatty acid production (Blomquist et al., 1994). The housefly microsomal and soluble FASs were purified to homogeneity (Gu et al., 1997) and the microsomal FAS was shown to preferentially use methylmalonyl-CoA in comparison to the soluble FAS. GC–MS analyses showed that the methyl-branching positions of the methyl-branched fatty acids of the housefly (Blomquist et al., 1994) were in positions consistent with them being the precursors of the methyl-branched hydrocarbons. The methylmalonyl-CoA unit that is the precursor to methyl-branched fatty acids and hydrocarbons arises from the carbon skeletons of valine and isoleucine, but not succinate (Dillwith et al., 1982). Propionate is also a precursor to methylmalonyl-CoA, and in the course of these studies, a novel pathway for propionate metabolism in insects was discovered. Many insect species, including the housefly, do not contain vitamin B12 (Wakayama et al., 1984), and therefore cannot catabolize propionate via methylmalonyl-CoA to succinate. Instead, as first demonstrated in the housefly (Dillwith et al., 1982), insects metabolize propionate to 3-hydroxypropionate and then to acetyl-CoA, with carbons 3 and 2 of propionate becoming carbons 1 and 2 of acetyl-CoA (Figure 8) (Halarnkar et al., 1986). 3.14.4.5.2. Pheromone biosynthesis in Drosophila The other major dipteran system in which pheromone production has been extensively studied is that of Drosophila. Jallon and Wicker-Thomas (2003) provide an excellent and detailed review of the chemistry, biochemistry, molecular biology, and genetics of pheromone production in Drosophila. Drosophila, as do Musca, use long chain, sexspecific hydrocarbons as close range and contact sex pheromones. Drosophila melanogaster Canton-S mature females have abundant (Z,Z)-7,11heptacosadiene, which is absent on males and was shown to effectively stimulate wing vibration

723

in conspecific males when applied to a dummy (Antony and Jallon, 1982; Ferveur and Sureau, 1996). In some Drosophila, 7-tricosene is involved in chemical communication. A series of studies established that the hydrocarbon pheromones are produced in the oenocytes (Ferveur et al., 1997), transported through the hemolymph via lipophorin (Pho et al., 1996), and then deposited on the cuticle. A series of experiments assaying the incorporation of various precursors into hydrocarbons established that Drosophila use an elongation–decarboxylation pathway for hydrocarbon production (Pennanec’h et al., 1997). An interesting question in Drosophila pheromone production is the origin of the 7,11 double bonds. Biochemical evidence suggested that either a D7-desaturase working on myristate, or a D9-desaturase working on palmitate give rise to the double bond in the 7-position, which could then be elongated to produce vaccenate (n-7, 18 : 1). Vaccenate could then be elongated and decarboxylated to produce 7-tricosene, and, with an additional desaturation step, to produce 7,11-27:2Hyd. Wicker-Thomas et al. (1997) isolated a cDNA encoding a desaturase (desat1) in D. melanogaster. The expressed desat1 protein is a D9-desaturase that preferentially used palmitate, resulting in n-7 fatty acids. A desat2, located close to desat1, appears to be responsible for the 5,9-dienes present in Tai females (Jallon and Wicker-Thomas, 2003). The identification of the desaturase genes responsible for pheromone production in fruit flies will greatly benefit from the characterization of the D. melanogaster genome. Understanding of the genetics combined with molecular biology undoubtedly will result in a more complete understanding of the mechanisms and genes involved and regulated in pheromone production in Drosophila. 3.14.4.6. Biosynthesis of Contact Pheromones in the German Cockroach

Upon antennal contact with a female, the male German cockroach rotates his body 180 and raises his wings, thus exposing specialized tergal glands that attract the female and place her into a precopulatory position (Nojima et al., 1999). The nonvolatile contact pheromone responsible for this behavior was identified as (3S,11S)-dimethylnonacosan-2one (Figure 1) (Nishida et al., 1974), and an alcohol (29-hydroxy-3S,11S-dimethylnonacosan-2-one) and an aldehyde (29-oxo-3,11-dimethylnonacosan-2one) derivatives with the same 3,11-dimethylketone skeleton (Nishida and Fukami, 1983). A fourth pheromone component, 3,11-dimethylheptacosan2-one, is less active than its C29 homolog (Schal et al., 1990b).

724 Biochemistry and Molecular Biology of Pheromone Production

The route of biosynthesis and its physiological regulation have been reviewed previously (Blomquist et al., 1993; Tillman et al., 1999; Schal et al., 2003). Central to investigations of the biosynthetic pathway was the observation that the major cuticular hydrocarbon in all life stages of the German cockroach is an isomeric mixture of 3,7-, 3,9- and 3,11-dimethylnonacosane (Jurenka et al., 1989). The presence of only the 3,11-isomer in the cuticular dimethyl ketone fraction and only in adult females prompted Jurenka et al. (1989) to propose that production of the pheromone might result from the sex-specific oxidation of its hydrocarbon analog only in adult females. This scheme follows the wellestablished conversion of hydrocarbons to methyl ketone and epoxide pheromones in the housefly (see above) (Blomquist et al., 1984b; Ahmad et al., 1987). This model has since been validated with several independent approaches. Biochemical studies on the biosynthesis of methyl-branched alkanes showed that the methyl branches are added during the early stages of chain elongation (Chase et al., 1990). Using carbon-13 labeling and NMR analyses, Chase et al. (1990) showed that carbons 1 and 2 of acetate are incorporated as the chain initiator, and that the carbon skeleton of propionate serves as the methyl branch donor (Figure 9). Further, propionate and succinate labeled methyl-branched hydrocarbons and the methyl ketone pheromone, as did the amino acids valine, isoleucine, and methionine, all of which can be metabolized to propionate. NMR studies confirmed that these substrates were metabolized to methylmalonyl-CoA for incorporation into the methyl branch unit of hydrocarbons (Chase et al., 1990), as in the housefly (Dillwith et al., 1982; Halarnkar et al., 1986), American cockroach (Halarnkar et al., 1985), cabbage looper moth (de Renobales and Blomquist, 1983), and the termite Zootermopsis (Chu and Blomquist, 1980). Methyl-branched fatty acids are intermediates in branched alkane biosynthesis (Juarez et al., 1992). Thus, [14C1]propionate labeled methyl-branched fatty acids of 16–20 carbons, but did not label straight-chain saturated and monounsaturated fatty acids (Chase et al., 1990). Chase et al. (1992) investigated the hypothesis that the 3,11-dimethyl ketone sex pheromone arises from the insertion of an oxygen into the preformed 3,11-dimethyl alkane. When high-specific activity, tritiated 3,11-dimethylnonacosane (mixture of stereoisomers), was topically applied on the cuticle of B. germanica females, it readily penetrated the cockroach and radioactivity from the alkane was detected in both 3,11-dimethylnonacosan-2-ol and

3,11-dimethylnonacosan-2-one. Likewise, when tritiated 3,11-dimethylnonacosan-2-ol was applied to the cuticle it was readily and highly efficiently converted to the corresponding methyl ketone pheromone. But, surprisingly, the dimethyl ketone pheromone was derived from the corresponding alcohol not only in females, as expected, but also in males. These results suggest that the sex pheromone of B. germanica arises via a female-specific hydroxylation of 3,11-dimethylnonacosane and a subsequent nonsex-specific oxidation, probably involving a polysubstrate monooxygenase system, to the (3S,11S)-dimethylnonacosan-2-one pheromone (Figure 9). Chase et al. (1992) also suggested that a similar hydroxylation and subsequent oxidation at the 29-position of 3,11-dimethylnonacosan-2-one might give rise to 29-hydroxy- and 29-oxo-(3,11)dimethylnonacosan-2-one, the other components of the contact pheromone blend, but this hypothesis has yet to be tested. It is quite likely, as well, that the same mechanism converts 3,11-dimethylheptacosane to the corresponding methyl ketone pheromone, and perhaps its 27-hydroxy- and 27-oxo- analogs. The contact sex pheromone of female B. germanica remains the only cockroach pheromone whose biosynthetic pathway has been investigated with radio- and stable-isotope tracers. 3.14.4.7. Biosythesis of the Honeybee Queen Pheromone

The queen substance used for ‘‘queen control’’ inside the nest is also the substance used by virgin queens to attract drones for mating. It is the best understood of the sexual pheromones of the social insects. Callow and Johnston (1960) and Barbier and Lederer (1960) identified ([E]-9-oxodec-2-enoic acid) (9-ODA) in queen mandibular glands. 9Hydroxy-2(E)-decenoic acid (9-HDA) is also present (Callow et al., 1964) and together both attract drones. Recent work (Keeling et al., 2003) identified a number of additional compounds that function synergistically with the 9-ODA and 9-HDA, making this the most complex pheromone blend known for any organism. In an elegant set of experiments, Plettner et al. (1996, 1998) elucidated the biosynthetic pathways for the honeybee queen mandibular pheromone (QMP) components 9-ODA and 9-HDA and compared their biosyntheses to that of worker produced 10-hydroxy-2(E)-decenoic acid and the corresponding diacid. Using carbon-13 and deuterated precursors, Plettner et al. (1996, 1998) demonstrated (1) the de novo synthesis of stearic acid in worker mandibular glands, (2) the hydroxylation of

Biochemistry and Molecular Biology of Pheromone Production

f0045

725

Figure 9 Proposed biosynthetic pathways for the major pheromone components of the German cockroach, Blattella germanica. The JH III regulated step appears to be the hydroxylation of the dimethylalkane.

726 Biochemistry and Molecular Biology of Pheromone Production

stearic acid at the n-(workers) and n-1 (queens) positions, (3) chain shortening through b-oxidation to the 10- and 8-carbon hydroxy acids, and (4) oxidation of n- and n-1 hydroxy groups to give diacids and 9-keto-2(E)-decenoic acid, respectively. Stearic acid was shown to be the main precursor of the pheromone molecules as it was converted to C10 hydroxy acids and diacids more efficiently than either 16 or 14 carbon fatty acids.

3.14.5. Endocrine Regulation of Pheromone Production 3.14.5.1. Barth’s Hypothesis

Juvenile hormone’s central role in mate-finding was recognized in 1965 when Barth proposed that neuroendocrine control of pheromone production would be common in insects with a long-lived adult stage and with multiple reproductive cycles interrupted by periods during which sexual receptivity and mating are not appropriate or not even possible anatomically. Cockroaches and beetles are quintessential examples of this life-history syndrome. Conversely, in insects that eclose with mature oocytes, and live for only a few days as adults, Barth (1965) predicted that pheromone signaling would be part of the adult metamorphic process and not subject to neuroendocrine control. The discovery of PBAN in moths (see Section 3.14.4.5.2) appeared in conflict with this hypothesis, but Barth’s model (Barth and Lester, 1973) clearly accounted for cases in moth species where adults feed and oocyte maturation requires the participation of JH or other neuroendocrine factors. Schal et al. (2003) proposed a reconsideration of the hypothesis, taking into account the coordination of reproductive developmental processes with mating-related events. Accordingly, in long-lived insects, such as cockroaches, pheromone production is expected to be synchronously regulated with other reproductive processes by the same hormone, usually JH. Cellular remodeling of the pheromone glands plays a prominent role in this group of insects, resulting in a slow stimulation of pheromone production. The cessation of pheromone production after mating is also slow, and precise control of pheromone signaling, therefore, is not at the level of pheromone production, but rather at the behavioral level through control of pheromone emission during calling. Conversely, in short-lived moths rapid modulation of rate-limiting enzymes in the pheromone biosynthetic pathway is much more prominent than developmental processes, and pheromone biosynthesis is turned on or off in coordination with activity cycles

(day versus night) and sexual receptivity (virgin versus mated). Control of sexual signaling occurs at the level of pheromone production as well as emission, but these two events are usually regulated by different factors. Thus, both groups of insects exhibit neuroendocrine control of pheromone production. In cockroaches, pheromone production is coordinated with the gonotrophic cycle and the major gonadotropic hormone – JH – has been recruited to control both by acting at several target tissues. In most moths, alternatively, reproduction and pheromone production are regulated by different hormones. But here also, the hormones that control pheromone production (e.g., PBAN) also affect other target tissues such as myotropins, melanization agents, and diapause and pupariation factors. An interesting departure from the moth model occurs in migratory moth species in which reproduction is delayed by migration (low levels of JH production and sexual inactivity), and pheromone production and its release are JH dependent (Cusson et al., 1994). All these observations are consistent with our interpretation of Barth’s model. The three hormones that regulate pheromone production in insects are shown in Figure 2 and Table 1. PBAN has been studied in female moths and alters enzyme activity through second messengers at one or more steps during or subsequent to fatty acid synthesis during pheromone production (Rafaeli and Jurenka, 2003). In contrast, 20E and JH induce or repress the synthesis of specific enzymes at the transcription level. The action of JH has been studied most thoroughly in the German cockroach and in bark beetles, and this work is discussed below. Ecdysteroid regulation of pheromone production occurs in Diptera, and has been most extensively studied in the housefly, M. domestica. 3.14.5.2. PBAN Regulation in Moths

3.14.5.2.1. PBAN Most female moths release sex pheromones in a typical calling behavior in which the pheromone gland is extruded to release pheromone during a particular time of the photoperiod. In most cases pheromone biosynthesis coincides with calling behavior and the synchronization of these events is achieved by neuroendocrine mechanisms present in the female that in turn are influenced by various environmental and physiological events such as temperature, photoperiod, host plants, mating, hormones, neurohormones, and neuromodulators. We now know that the main neuroendocrine mechanism that regulates pheromone production in moths is pheromone biosynthesis activating neuropeptide (PBAN).

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Table 1 Amino acid sequences of the pyrokinin/PBAN family of peptides; the FXPRLamide motif is shown in bold (X standing for S, T, G, or V) Function

Species

Peptide sequence

Reference

PBAN

Helicoverpa zea Helicoverpa assulta Bombyx mori Lymantria dispar Agrotis ipsilon Mamestra brassicae Spodoptera littoralis Bombyx mori Helicoverpa zea Helicoverpa assulta Agrotis ipsilon Mamestra brassicae Spodoptera littoralis Bombyx mori Helicoverpa zea Helicoverpa assulta Agrotis ipsilon Mamestra brassicae Pseudaletia separata Spodoptera littoralis Bombyx mori Helicoverpa zea Helicoverpa assulta Agrotis ipsilon Mamestra brassicae Spodoptera littoralis Bombyx mori Helicoverpa zea Helicoverpa assulta Agrotis ipsilon Spodoptera littoralis Leucophaea maderae Locusta migratoria

LSDDMPATPADQEMYRQDPEQIDSRTKYFSPRLamidea LSDDMPATPADQEMYRQDPEQIDSRTKYFSPRLamideb LSEDMPATPADQEMYQPDPEEMESRTRYFSPRLamidea LADDMPATMADQEVYRPEPEQIDSRNKYFSPRLamidea LADDTPATPADQEMYRPDPEQIDSRTKYFSPRLamideb LADDMPATPADQEMYRPDPEQIDSRTKYFSPRLamideb LADDMPATPADQELYRPDPDQIDSRTKYFSPRLamideb a IIFTPKLamideb a VIFTPKLamideb a VIFTPKLamideb a VIFTPKLamideb a VIFTPKLamideb a VIFTPKLamideb b SVAKPQTHESLEFIPRLamideb b SLAYDDKSFENVEFTPRLamideb b SLAYDDKSFENVEFTPRLamideb b SLSYEDKMFDNVEFTPRLamideb b SLAYDDKVFENVEFTPRLamideb b KLSYDDKVFENVEFTPRLamideb b SLAYDDKVFENVEFTPRLamideb g TMSFSPRLamideb g TMNFSPRLamideb g TMNFSPRLamideb g TMNFSPRLamideb g TMNFSPRLamideb g TMNFSPRLamideb TDMKDESDRGAHSERGALCFGPRLamidea NDVKDGAASGAHSDRLGLWFGPRLamideb NDVKDGAASGAHSDRLGLWFGPRLamideb NDVKDGGADRGAHSDRGGMWFGPRIamideb NEIKDGGSDRGAHSDRAGLWFGPRLamideb ETSFTPRLamidea (I) pEDSGDGWPQQPFVPRLamidea (II) pESVPTFTPRLamidea (I) GAVPAAQFSPRLamidea (II) EGDFTPRLamidea (III) RQQPFVPRLamidea (IV) RLHQNGMPFSPRLamidea MT1 GAAPAAQFSPRLamidea MT2 TSSLFPHPRLamidea PK1 HTAGFIPRLamidea PK2 SPPFAPRLamidea PK3 LVPFRPRLamidea PK4 DHLPHDVYSPRLamidea PK5 GGGGSGETSGMWFGPRLamidea PK6 SESEVPGMWFGPRLamidea PK4 DHLSHDVYSPRLamidea CAP2b-3 TGPSASSGLWFGPRLamideb

Raina et al. (1989) Choi et al. (1998) Kitamura et al. (1989) Masler et al. (1994) Duportets et al. (1999) Jacquin-Joly et al. (1998) Iglesias et al. (2002) Kawano et al. (1992) Ma et al. (1994) Choi et al. (1998) Duportets et al. (1999) Jacquin-Joly et al. (1998) Iglesias et al. (2002) Kawano et al. (1992) Ma et al. (1994) Choi et al. (1998) Duportets et al. (1999) Jacquin-Joly et al. (1998) Matsumoto et al. (1992) Iglesias et al. (2002) Kawano et al. (1992) Ma et al. (1994) Choi et al. (1998) Duportets et al. (1999) Jacquin-Joly et al. (1998) Iglesias et al. (2002) Imai et al. (1991) Ma et al. (1994) Choi et al. (1998) Duportets et al. (1999) Iglesias et al. (2002) Holman et al. (1986) Schoofs et al. (1991) Schoofs et al. (1993) Schoofs et al. (1990a) Schoofs et al. (1990b) Schoofs et al. (1992) Schoofs et al. (1992) Veelaert et al. (1997) Veelaert et al. (1997) Predel et al. (1997) Predel et al. (1997) Predel et al. (1999) Predel et al. (1999) Predel et al. (1999) Predel and Eckert (2000) Predel and Eckert (2000) Choi et al. (2001)

PK-2 SVPFKPRLamideb ETH-1 DDSSPGFFLKITKNVPRLamideb hug g pELQSNGIPAYRVRTPRLamideb DFAFSPRLamidea

Choi et al. (2001) Park et al. (1999) Meng et al. (2002) Torfs et al. (2001)

ADFAFNPRLamidea

Torfs et al. (2001)

Pheromontropic peptides

Diapause hormone

Pyrokinins

Myotropins

Locusta migratoria

Schistocerca gregaria Periplaneta americana

Periplaneta fuliginosa Drosophila melanogaster

Penaeus vannamei

(Crustacea)

a b

Identified from the amino acid sequence of a purified peptide. Deduced from the cloned gene sequence.

728 Biochemistry and Molecular Biology of Pheromone Production

The neuropeptide PBAN was localized to the subesophageal ganglion (SEG) (Raina and Menn, 1987), which facilitated purification and sequencing (Kitamura et al., 1989; Raina et al., 1989). The first PBAN identified had 33 amino acids with a C-terminal amidation and the core sequence FXPRLamide (where X represents S, T, G, or V) is required for activity, which places PBAN in a family of peptides with the C-terminal FXPRLamide motif (Table 1). The first member of this family to be identified was called leucopyrokinin based on its ability to stimulate hindgut contraction in the cockroach Leucophaea maderae (Holman et al., 1986). Additional functions for this family include induction of embryonic diapause in B. mori (Imai et al., 1991), induction of melanization in lepidopteran larvae (Matsumoto et al., 1990), and acceleration of puparium formation in several flies (Zdarek et al., 1998). In addition it was determined that the white shrimp, Penaeus vannamei, has two peptides that can induce myotropic activity (Torfs et al., 2001). These results demonstrate the ubiquity and multifunctional nature of this family of peptides. Localization of PBAN-like immunoreactivity in the central nervous system of adult moths indicated that several neurons in the SEG contain PBAN-like activity. These were found as clusters along the ventral midline, one each in the presumptive mandibular, maxillary, and labial neuromeres (Kingan et al., 1992). All three groups of neurons have axons that project into the corpus cardiacum (CC) (Kingan et al., 1992; Davis et al., 1996). Two pairs of maxillary neurons send processes within the SEG that include posterior projections into the paired ventral nerve cord (VNC) and travel its entire length to terminate in the terminal abdominal ganglion (TAG). Arborizations arising from these paired projections were found in each segmental ganglion (Davis et al., 1996). In addition the segmental ganglia have neurons that contain PBAN-like activity and these neurons can release peptides into the hemolymph (Ma and Roelofs, 1995c; Davis et al., 1996; Ma et al., 1996). 3.14.5.2.2. Molecular genetics of PBAN The gene encoding PBAN was first characterized from H. zea and B. mori (Imai et al., 1991; Davis et al., 1992; Kawano et al., 1992; Sato et al., 1993; Ma et al., 1994). The full-length cDNA was found to encode PBAN plus four additional peptide domains with a common C-terminal FXPRL sequence motif including that of the diapause hormone of B. mori. Three additional peptides with the common Ctermini and sequence homology to those of H. zea and B. mori have been deduced from cDNA isolated

from pheromone glands of Mamestra brassicae (Jacquin-Joly and Descoins, 1996), H. assulta (Choi et al., 1998), Agrotis ipsilon (Duportets et al., 1999), Spodoptera littoralis (Iglesias et al., 2002), H. armigera (Zhang et al., 2001), and Adoxophyes sp. (Lee et al., 2001). The posttranslational processed peptides can be found in the SEG (Sato et al., 1993; Ma et al., 1996). The matrix-assisted laser desorption ionization (MALDI) MS data indicated that PBAN was found to a greater extent in the mandibular and maxillary clusters than in the labial cluster (Ma et al., 2000). The other neuropeptides were found in all clusters. In addition some larger peptide fragments were found indicating alternative processing of the precursor protein (Ma et al., 2000). 3.14.5.2.3. PBAN mode of action Through the development of a sensitive in vitro bioassay, studies on H. armigera and H. zea demonstrated that brain extracts and synthetic H. zea PBAN could stimulate the production of the main pheromone component (Soroker and Rafaeli, 1989; Rafaeli et al., 1990, 1991, 1993; Rafaeli, 1994; Rafaeli and Gileadi, 1996). The response obtained was specific to the pheromone gland and independent of other abdominal tissues (Rafaeli, 1994; Rafaeli et al., 1997b). Evidence has since accumulated regarding several other species of moths including B. mori, Spodoptera litura, O. nubilalis, Plodia interpunctella, and Thaumetopoea pityocampa (Arima et al., 1991; Fo´ nagy et al., 1992; Fabria`s et al., 1995; Ma and Roelofs, 1995b; Rafaeli and Gileadi, 1995a; Jurenka, 1996). In addition, functional and viable pheromone gland cell-clusters were obtained from the intersegmental membrane of B. mori using papain enzymatic digestion (Fo´ nagy et al., 2000). The signal transduction events that occur after PBAN binds to a receptor have been studied in several model moth species (Figure 10). The main difference found between these species so far is whether or not 30 ,50 ,cyclic-AMP (cAMP) is used as a second messenger. In the case of the heliothines and several others, cAMP is a second messenger. Alternatively, cAMP is thought not to act in pheromone gland cells of B. mori and O. nubilalis (Fo´ nagy et al., 1992; Ma and Roelofs, 1995b). Instead, in these insects, it is thought that an increase in cytosolic calcium directly activates downstream events leading to stimulation of the biosynthetic pathway. Extracellular calcium is essential for pheromonotropic activity in all moths studied to date (Fo´ nagy et al., 1992, 1999; Ma and Roelofs, 1995a; Matsumoto et al., 1995). It is suggested that free calcium,

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Figure 10 Proposed signal transduction mechanisms that stimulate the pheromone biosynthetic pathway in Helicoverpa zea and other heliothines as compared with that in Bombyx mori. It is proposed that PBAN binds to a receptor present in the cell membrane. Binding to the receptor somehow induces a receptor-activated calcium channel to open causing an influx of extracellular calcium. This calcium binds to calmodulin and in the case of B. mori will directly stimulate a phosphatase that will dephosphorylate and activate a reductase in the biosynthetic pathway. This activated reductase will then produce the pheromone bombykol. In H. zea and other heliothines like H. armigera the calcium-calmodulin will activate adenylate cyclase to produce cAMP that will then act through kinases and/or phosphatases to stimulate acetyl-CoA carboxylase in the biosynthetic pathway. (Reprinted with permission from Rafaeli, A., Jurenka, R.A., 2003. PBAN regulation of pheromone biosynthesis in female moths. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Pheromone Biochemistry and Molecular Biology. Elsevier, San Diego, CA, pp. 107–136; ß Elsevier.)

entering the cell, binds to calmodulin to form a complex thereby activating adenylate cyclase and/or phosphoprotein phosphatases (Figure 10). Calmodulin was characterized from pheromone glands of B. mori (Iwanaga et al., 1998) and was shown to have an identical amino acid sequence to Drosophila calmodulin (Smith et al., 1987). In B. mori, it is suggested that the Ca2þ/calmodulin complex directly or indirectly activates a phosphoprotein phosphatase (Matsumoto et al., 1995). This phosphatase will then activate an acyl-CoA reductase in the biosynthetic pathway. Two genes encoding calcineurin heterosubunits were identified from the pheromone gland of B. mori and were found to be homologous to the catalytic subunit and regulatory subunits of other animal calcineurins (Yoshiga et al., 2002). The calcineurin complex will apparently dephosphorylate an acyl-CoA reductase, which catalyzes the formation of bombykol in B. mori. 3.14.5.2.4. Enzymes affected in the pheromone biosynthetic pathway PBAN has been shown to stimulate the reductase that converts an acyl-CoA

to an alcohol precursor (Figure 10) in several moths including B. mori (Arima et al., 1991; Ozawa et al., 1993), T. pityocampa (Gosalbo et al., 1994), S. littoralis (Martinez et al., 1990; Fabria`s et al., 1994), and M. sexta (Fang et al., 1995b; Tumlinson et al., 1997). In A. velutinana (Tang et al., 1989), H. zea (Jurenka et al., 1991a), Cadra cautella, S. exigua (Jurenka, 1997) and M. brassicae (Jacquin et al., 1994), it was demonstrated that PBAN controls pheromone biosynthesis by regulating a step during or prior to fatty acid biosynthesis (Figure 10). Circumstantial evidence in A. segetum (Zhu et al., 1995) and H. armigera (Rafaeli et al., 1990) also points to the regulation of fatty acid synthesis by PBAN. In one study using the moth Sesamia nonagrioides it was shown that the acetyltransferase enzyme might be regulated by PBAN (Mas et al., 2000). There appears to be no particular pattern as to which enzyme within the pheromone biosynthetic pathway will be regulated by PBAN. However, in the majority of moths studied it is either the reductase or fatty acid synthesis that is stimulated.

730 Biochemistry and Molecular Biology of Pheromone Production

Several families of moths utilize hydrocarbons and/ or their epoxides as sex pheromones (Millar, 2000). It is thought that PBAN does not regulate the production of hydrocarbon sex pheromones as demonstrated in Scoliopteryx libatrix (Subchev and Jurenka, 2001). However, PBAN is probably regulating the production of epoxide sex pheromones. This was demonstrated in Ascotis selenaria cretacea where decapitation resulted in pheromone decline and it could be restored by injecting PBAN (Miyamoto et al., 1999). Decapitation also decreases the epoxide pheromone titer in the gypsy moth, L. dispar, and injection of PBAN can restore pheromone production (Thyagaraja and Raina, 1994). However, decapitation did not decrease the levels of the hydrocarbon precursor in the gypsy moth (Jurenka, unpublished data). These findings indicate that PBAN may regulate the epoxidation step in those moths that utilize epoxide pheromones but not the production of the alkene precursor or alkene pheromones. 3.14.5.2.5. Mediators and inhibitors of PBAN action Juvenile hormones play an important role in reproductive development of many moth species (see Chapters 3.7 and 3.9). Although JH probably does not regulate pheromone biosynthesis directly it has been shown to be involved in the release of PBAN in the migratory moths Pseudaletia unipuncta and A. ipsilon (Cusson and McNeil, 1989; Gadenne, 1993; Cusson et al., 1994; Picimbon et al., 1995). In addition, JH has been shown to prime the pheromone glands in pharate adults of the nonmigratory moth H. armigera (Fan et al., 1999). JH II, in an in vitro assay, primed pheromone glands of pharate adults to respond to PBAN and induced earlier pheromone production by intact newly emerged females (Fan et al., 1999). This induction could be mediated by JH upregulation of a putative PBAN receptor in pharate adults (Rafaeli et al., 2003). The corpus bursae has been implicated in the mediation of PBAN stimulation of pheromone biosynthesis in some tortricids (Jurenka et al., 1991c). A peptide was partially purified from corpus bursae that could stimulate pheromone production (Fabria`s et al., 1992). To date a bursal factor has only been demonstrated in A. velutinana and the related tortricids C. fumiferana and C. rosaceana (Delisle et al., 1999). The role of the nervous system in pheromone biosynthesis in moths is not clearly understood. In several moths including L. dispar (Tang et al., 1987; Thyagaraja and Raina, 1994), H. virescens (Christensen et al., 1991), S. littoralis (Marco et al., 1996), and M. brassicae (Iglesias et al., 1998) an

intact VNC was reported as necessary for pheromone biosynthesis. Christensen and coworkers (Christensen et al., 1991, 1992, 1994; Christensen and Hildebrand, 1995) proposed that the neurotransmitter octopamine might be involved as an intermediate messenger during the stimulation of sex pheromone production in H. virescens. These workers suggested that octopamine was involved in the regulation of pheromone production and that PBAN’s role lies in the stimulation of octopamine release at nerve endings. However, contradicting results concerning VNC transection and octopamine-stimulated pheromone production were reported in the same species as well as in other moth species (Jurenka et al., 1991c; Ramaswamy et al., 1995; Rafaeli and Gileadi, 1996; Park and Ramaswamy, 1998; Delisle et al., 1999). A modulatory role for octopamine was suggested by research conducted on H. armigera (Rafaeli and Gileadi, 1995b; Rafaeli et al., 1997a, 1999). Octopamine and several octopaminergic analogs inhibited pheromone production in studies using both in vitro and in vivo bioassays in two species of moths (Rafaeli and Gileadi, 1995b, 1996; Rafaeli et al., 1997, 1999; Hirashima et al., 2001). The role of the VNC in pheromone production still requires clarification. It has been suggested, that in some moth species (S. littoralis) both humoral and neural regulation occurs (Marco et al., 1996). Given the diversity of moths it may not be surprising to find several mechanisms regulating pheromone biosynthesis. 3.14.5.3. JH Regulation in Beetles

3.14.5.3.1. General In the Coleoptera, pheromone production or release is controlled by JH III (review: Tillman et al., 1999). JH, or JH analogs, stimulate pheromone production in T. molitor (Menon, 1970), T. castaneum (both Tenebrionidae), some members of the Cucujidae (review: Plarre and Vanderwel, 1999), A. grandis (Curculionidae) (Wiygul et al., 1990), and various Scolytidae (see Section 3.14.5.3.2). Various factors may work upstream as cues to stimulate JH biosynthesis. Feeding stimulates pheromone production or release in many species (Vanderwel and Oehlschlager, 1987). Sexual maturity and population density may also be important, e.g., for some Scolytidae (Byers, 1983) and Cucujidae (Plarre and Vanderwel, 1999). The effect of population density may be mediated by sensitivity to pheromone concentrations. Antennectomy can raise pheromone biosynthetic rates in A. grandis (Dickens et al., 1988) and L. decemlineata (Dickens et al., 2002), suggesting that detection of pheromone components may inhibit their production. Various combinations of physiological and

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environmental factors therefore regulate JH titers, which in turn stimulate pheromone biosynthesis and/or release.

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3.14.5.3.2. JH regulation in bark beetles (Scolytidae) Aggregation pheromone biosynthesis in bark beetles generally begins shortly after the beetle arrives at a new host tree (Wood, 1982; Seybold et al., 2000). Unfed beetles can be artificially stimulated to produce pheromones by treatment with JH III. For example, males are the pioneers of Ips spp., and starved male I. paraconfusus and I. pini can be induced to synthesize the pheromone components ipsenol and ipsdienol if JH III or a JH analog is applied topically (Bordon et al., 1969; Chen et al., 1988; Ivarsson and Birgersson, 1995; Tillman et al., 1998). Similarly, starved Dendroctonus spp. can be induced to synthesize pheromone components following treatment with JH (Bridges, 1982; Conn et al., 1984). Feeding apparently stimulates the synthesis of JH III in the CA (Tillman et al., 1998), resulting in elevated JH titers that trigger pheromone biosynthesis in midgut cells (Hall et al., 2002a, 2002b). Pheromone biosynthesis requires a shift in metabolic priorities, particularly for those beetles that produce large quantities of pheromone. JH must actuate this shift, and since pheromone components are synthesized de novo (Seybold et al., 1995b), induction of the pheromone biosynthetic pathway involves elevated expression of at least some related genes. This was first demonstrated in I. paraconfusus, where JH-treated male beetles have increased HMG-R mRNA levels compared to controls (Tittiger et al., 1999). Male I. pini and D. jeffreyi HMG-R mRNA levels also rise in response to topical JH III applications (Figure 11) (Tittiger et al., 2003; Tillman, Lu, Tittiger, Blomquist, and Seybold, unpublished data). In all species studied, the response is dose and time dependent and correlates with pheromone component biosynthesis. This is consistent with the hypothesis that JH controls HMG-R, which functions as a regulator of the mevalonate (pheromone biosynthetic) pathway. HMG-S mRNA levels in male D. jeffreyi also respond to topical JH applications similarly to HMG-R, though at a more modest level (Figure 11) (Tittiger et al., 2000). In I. pini, mRNA for a GPPS gene is also elevated in JH III-treated beetles (Gilg-Young, Welch, Tittiger, and Blomquist, unpublished data). This enzyme diverts carbon from the normal mevalonate pathway into the pheromone biosynthetic pathway, and is thus also a likely regulatory step. The HMG-S and HMG-R studies in D. jeffreyi show how putative regulatory enzymes are

Figure 11 Regulation of HMG-R and HMG-S expression in male Dendroctonus jeffreyi by JH III. The time course (a, b) and dose response (c) for each gene in mature (emerged) males was investigated by Northern blotting. All values are relative to starved, untreated males. Each point represents the mean þ/ standard error of three replicates, five isolated thoraces/sample. (Reprinted with permission from Tittiger, C., Barkawi, L.S., Bengoa, C.S., Blomquist, G.J., Seybold, S.J., 2003. Structure and juvenile hormone-mediated regulation of the HMG-CoA reductase gene from the Jeffrey pine beetle, Dendroctonus jeffreyi. Mol. Cell. Endocrinol. 199, 11–21; ß Elsevier.)

controlled at the transcriptional level by JH III (though posttranscriptional regulation is probably also important; see below). Their coordinate induction implies that other mevalonate/pheromone biosynthetic genes may also be stimulated by JH III. This question has been addressed mostly in I. pini; indeed, more is known about pheromone biosynthetic gene regulation in this insect than in any other beetle, due in part to microarray-based expression profiling. Microarray analyses of male and female JH-treated and untreated I. pini give an idea of the profound change that male midgut cells undergo in order to synthesize pheromone, with numerous genes being upregulated or downregulated

732 Biochemistry and Molecular Biology of Pheromone Production

following JH III treatment (Keeling and Tittiger, unpublished data). All identified mevalonate pathway genes in a recent expressed sequence tags (ESTs) project (BeetleBase, 2003; Eigenheer et al., 2003) have elevated expression levels compared to controls (Keeling, Bearfield, Blomquist, and Tittiger, unpublished data). These include enzymes at predicted control points (HMG-R and GPPS) as well as others that would not be expected to have a strong regulatory role (e.g., isopentenyl diphosphate (IPP) isomerase). Thus, induction of identified pheromone-biosynthetic genes is apparently coordinate. By extension, genes encoding unidentified enzymes in the pheromone biosynthetic pathway are also likely JH regulated. The putative pheromone precursor, ipsdienone, was recovered from thoracic sections of JH-treated I. paraconfusus (Ivarsson et al., 1998), and a male-specific myrcene synthase activity is also elevated in tissues of JH III-treated I. pini (Martin et al., 2003). These data provide biochemical evidence that JH III affects enzyme activities converting geranyl diphosphate to ipsdienol, with myrcene and ipsdienone as probable intermediates. Confirmation that the corresponding genes are also JH regulated awaits their identification, and characterization of the relevant enzymes. In the studies mentioned above, the amount of JH III penetrating the cuticle and acting on the midguts is unknown. A more biologically relevant question is whether feeding similarly stimulates pheromone biosynthetic gene expression. Recent real-time polymerase chain reaction (PCR) studies of RNA recovered from midguts from fed male and female I. pini confirm that HMG-R and HMG-S are induced in both sexes by feeding to levels similar to those observed in JH III-treated insects (Keeling and Tittiger, unpublished data). This is curious, because female midguts do not synthesize monoterpenoid pheromones, and female JH III titers do not naturally rise upon feeding (Tillman et al., 1998). Other factors therefore must cause elevated HMG-R and HMG-S expression in female midguts. These experiments underscore the complexity of gene regulation in bark beetles, and remind us that mevalonate pathway genes almost certainly respond to other, possibly nonpheromone-related factors. Whatever the case, feeding clearly induces expression of certain genes in both male and female tissues, with males having an additional response characterized by increased rates of pheromone biosynthesis. Juvenile hormone may act as a both trigger and stimulus for pheromone production. The basal HMG-R and HMG-S mRNA levels are from fivefold to eightfold higher in the midguts of males than of females, implying a greater capacity of

mevalonate pathway flux in male cells compared to female cells (Keeling and Tittiger, unpublished data). Also, female GPPS mRNA levels, which are not clearly induced by feeding in female I. pini midguts, are approximately 16-fold lower than those in male midguts, and GPPS is induced by feeding and JH III treatment in males (Gilg-Young, Welch, Keeling, Bearfield, Blomquist, and Tittiger, unpublished data). This suggests that male midguts may be primed for pheromone biosynthesis, possibly due to an earlier, developmental cue, even before JH III triggers the process, and that pioneer beetles are ready to begin pheromone production when they arrive at a host tree. Subsequent feeding-induced JH III production thus both activates existing components of the pathway and stimulates continued production of pheromone pathway enzymes. Such priming might avoid a potentially dangerous ‘‘waiting’’ period, during which the pheromone-biosynthetic pathway is being stimulated, which would delay the arrival of other beetles to assist colonizing the host tree. JH appears to be necessary for pheromone production, but it is not always sufficient. Similar gene responses to topically applied JH in male and female I. pini belie very different metabolic responses (Tillman, Lu, Seybold, Keeling, Bearfield, Blomquist, and Tittiger, unpublished data), suggesting a control mechanism for I. pini that is more complex than having JH III simply stimulate expression of pheromone biosynthetic genes. Other factors may be involved. For example, while all mevalonate pathway genes are induced in male I. pini, only genes corresponding to early steps (e.g., thiolase, HMG-R, HMG-S, IPP isomerase) are induced by JH III in females, while those for latter steps (GPPS, farnesyl diphosphate synthase (FPPS)) are not (Keeling et al., 2004). Similarly, JH is clearly not sufficient to stimulate pheromone production in I. paraconfusus (Tillman, Lu, Tittiger, Blomquist, and Seybold, unpublished data). JH III-treated male I. paraconfusus have elevated HMG-R mRNA levels (the expression of other genes has not been studied in this insect), but neither HMG-R enzyme activity nor monoterpenoid pheromone component levels are significantly increased compared to controls. In contrast, male I. paraconfusus that have been allowed to feed on pine phloem have elevated HMG-R activity and, of course, elevated pheromone component levels. Decapitation studies confirm that an unknown ancillary hormone(s) is required in addition to JH III for pheromone biosynthesis (Lu, Blomquist, and Seybold, unpublished data). Presumably, one effect of this factor is in regulating the posttranscriptional activation of

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HMG-R. Posttranscriptional regulation of pheromone biosynthetic activity has been demonstrated in other insect systems (e.g., PBAN-regulated activity of bombykol in silkmoths) (Moto et al., 2003), and multilevel control of HMG-R activity is well documented (review: Hampton et al., 1996). Future studies that concentrate on the effects of JH III on protein levels and enzyme activities should provide exciting new information of how JH III regulates the pheromone-biosynthetic machinery in bark beetles.

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3.14.5.4. 20-Hydroxyecdysone Regulation of Pheromone Production in the Housefly

3.14.5.4.1. Ecdysteroid regulation Newly emerged female houseflies do not have detectable amounts of any C23 sex pheromone components. Sex pheromone production correlates with ovarian development and vitellogenesis. The C23 sex pheromone components first appear when ovaries mature to the early vitellogenic stages (Figure 12) and increase in amount until stages 9 and 10 (mature egg) (Dillwith et al., 1983; Mpuru et al., 2001). Females

Figure 12 Effect of ovarian development (a, b), ovariectomy and allatectomy (c, d, e), and treatment with 20-hydroxyecdysone (20E) on ovariectomized females (f, g) and males (h, i) on (Z )-9-tricosene production. The results show that ovariectomy abolishes pheromone production, whereas treatment with 20E induces pheromone production in ovariectomized females and in males. (Reprinted with permission from Blomquist, G.J., 2003. Biosynthesis and ecdysteroid regulation of housefly sex pheromone production. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Pheromone Biochemistry and Molecular Biology. Elsevier, New York, pp. 131–252; ß Elsevier.)

734 Biochemistry and Molecular Biology of Pheromone Production

ovariectomized within 6 h of adult emergence do not produce any of the C23 sex pheromone components, whereas control and allatectomized females produced abundant amounts of (Z)-9-tricosene (Figure 12c–e). Ovariectomized insects that received ovary implants produced sex pheromone components in direct proportion to ovarian maturation. These data demonstrate that a hormone from the maturing ovary induced sex pheromone production (Blomquist et al., 1993, 1998). Juvenile hormone regulates both vitellogenesis and pheromone production in some insect species (Tillman et al., 1999). In some Diptera, including the housefly, ovarian-produced ecdysteroids are involved in regulating vitellogenesis (Hagedorn, 1985; Adams et al., 1997) at the transcriptional level (Martin et al., 2001). Therefore, since ovariectomy abolished sex pheromone production whereas allatectomy (which abolishes JH production) had no effect on pheromone production (Blomquist et al., 1992), it was hypothesized that an ecdysteroid, and not JH, regulated sex pheromone production in the housefly. Injection of 20E at doses as low as 0.5 ng every 6 h induced sex pheromone production in ovariectomized houseflies in a time- and dosedependent manner (Figure 12f and g) (Adams et al., 1984a, 1984b, 1995). Multiple injections of 20E into ovariectomized insects over several days resulted in as much 23 : 1 produced as in intact control females. Application of JH or JH analogs alone or in combination with ecdysteroids had no effect on pheromone production (Blomquist et al., 1992). 3.14.5.4.2. Induction of female sex pheromone production in male houseflies Male houseflies normally produce no detectable C23 sex pheromone components, but do produce the same C27 and longer alkenes as previtellogenic females. Implantation of ovaries into male houseflies resulted in a change in the chain length specificity of the alkenes such that (Z)-9-tricosene became a major component (Blomquist et al., 1984a, 1987). Likewise, injection of 20E into males induces sex pheromone production in a dose-dependent manner (Figure 12h and i). Thus, males possess the biosynthetic capability to produce sex pheromone, but normally do not produce the 20E necessary to induce sex pheromone production. This makes male houseflies a very convenient model to study the regulation of sex pheromone production, circumventing the need to ovariectomize a large number of female insects. 3.14.5.4.3. Ecdysteroids affect fatty acyl-CoA elongation enzymes There are two likely possibilities to account for the change in the chain length

of the alkenes synthesized by the female housefly in the production of (Z)-9-trecosene. They are: (1) the chain-length specificity of the reductive conversion of acyl-CoAs to alkenes is altered such that 24:1CoA becomes an efficient substrate, or (2) there is a change in the chain-length specificity of the fatty acyl-CoA elongation enzymes such that 24:1-CoA is not efficiently elongated, resulting in an accumulation of 24:1-CoA. To determine which enzyme activities are affected by 20E to regulate the chain length of the alkenes, experiments were performed to examine the chain-length specificity of the fatty acyl-CoA reductive conversion of acyl-CoAs to alkenes and elongation. Microsomal preparations from both males and females of all ages examined readily converted 24:1-CoA and the 24:1 aldehyde to (Z)-9-tricosene, indicating that 20E was not acting on this activity (Tillman-Wall et al., 1992; Reed et al., 1995). In contrast, microsomes from day-4 females (high ecdysteroid titer and production of (Z)-9 tricosene) did not elongate either 18:1-CoA or 24:1-CoA beyond 24 carbons, while microsomes from day-4 males or day-1 females (both of which produce alkenes of 27:1 and longer) readily elongated both 18:1-CoA and 24:1-CoA to 28:1-CoA (TillmanWall et al., 1992; Blomquist et al., 1995). Thus, 20E appears to regulate the fatty acyl-CoA elongases and not the enzymatic steps in the conversion of acyl-CoA to hydrocarbon. 3.14.5.4.4. Transport of pheromone The role of hemolymph in transporting hydrocarbons and hydrocarbon pheromones has only recently become fully appreciated. Older models of hydrocarbon formation showed epidermal-related cells (oenocytes) synthesizing and transporting hydrocarbons directly to the surface of the insect (Hadley, 1984). In the housefly, the role of hemolymph is most clearly seen when (Z)-9-tricosene production is initiated. (Z)-9-Tricosene first accumulates in the hemolymph, and then, after a number of hours, is observed on the surface of the insect. Modeling of the process (Mpuru et al., 2001) showed that the delay is surprisingly long; a period of more than 24 h is necessary for transport from site of synthesis to deposition on the surface of the insect. In sexually mature females, (Z)-9-tricosene comprised a relatively large fraction of the hydrocarbon of the epicuticle and the hemolymph, but much smaller percentages of the hydrocabons in other tissues, including the ovaries. It appears that certain hydrocarbons were selectively partitioned to certain tissues such as the ovaries, from which pheromone was relatively excluded (Schal et al., 2001). Both KBr gradient ultracentrifugation and specific

Biochemistry and Molecular Biology of Pheromone Production

immunoprecipitation showed that over 90% of the hemolymph hydrocarbon was associated with a high-density lipophorin (see Chapter 4.6). Lipophorin was composed of two aproproteins under denaturing conditions: apolipophorin I (240 kDa) and apolipophorin II (85 kDa) (Schal et al., 2001). The data suggest that lipophorin may play an important role in an active mechanism that selectively delivers specific hydrocarbons to specific sites. 3.14.5.5. Regulation of Pheromone Production in Cockroaches

3.14.5.5.1. Development and cellular plasticity of pheromone glands In cockroaches, in striking contrast to many moths, pheromone glands acquire functional competence during an imaginal maturation period, and developmental regulation involves factors that also control adult reproductive readiness. Also, because reproduction in cockroaches is interrupted by periods of sexual inactivity (i.e., gestation), developmental regulation of the sex pheromone gland can result in alternating cycles of acquisition and subsequent waning of competence through maturation and retrogression, respectively, of cellular machinery. Consequently, in female cockroaches pheromone production is controlled by cyclic maturational changes in the gland in relation to the ovarian cycle. Best exemplifying this phenomenon are the tergal and sternal glands of N. cinerea and B. germanica males. Both species possess class-3 glandular units, composed of two cells – a secretory cell and a duct cell (Quennedey, 1998). But after apolysis and before the imaginal molt the immature gland contains four concentric cells, including in addition to the two adult cells an enveloping cell and a ciliary cell (Sreng and Quennedey, 1976; Sreng, 1998). During several days after the adult molt the gland matures, in part by undergoing apoptosis (programmed cell death). The ciliary cell gives rise to a part of the microvillar end-apparatus, then dies, whereas the enveloping cell forms an upper portion of the duct, then it too dies (Sreng, 1998). In concert, before day 5 the immature sternal glands of N. cinerea males produce little pheromone, but after day 5 their pheromone content increases significantly (Sreng et al., 1999). Decapitation or allatectomy of N. cinerea males completely blocked the apoptotic process, while JH III treatment restored apoptosis (Sreng et al., 1999). Brain extracts or synthetic moth PBAN failed to restore gland differentiation or stimulate pheromone production. Female B. germanica employ similar class-3 glands to produce a volatile sex pheromone that is yet to be identified. Ultrastructural, behavioral, and

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electrophysiological studies have shown that, as in males, pheromone gland cells mature as the female sexually matures (Abed et al., 1993b; Liang and Schal, 1993; Tokro et al., 1993; Schal et al., 1996). The secretory cells of newly formed glands in the imaginal female are small and they contain little pheromone (determined by behavioral and EAG assays). As the female sexually matures, the size of pheromone-secreting cells increases, as does its pheromone content (Liang and Schal, 1993). The mature pheromone gland then undergoes cycles of cellular hypertrophy and retrogression in relation to the JH III titer in successive reproductive cycles. The gland becomes atrophied and its pheromone content declines during gestation, but as a new vitellogenic cycle begins after the egg case is deposited, the pheromone gland undergoes rapid regrowth and proliferation of cellular organelles and an increase in its pheromone content. Although this pattern corresponds well with the JH III titer in the hemolymph (Liang and Schal, 1993; Schal et al., 1996), no experimental manipulations of hormone titers have been conducted to verify the hypothesis that JH III controls the cellular plasticity of the pheromone gland. 3.14.5.5.2. Pheromone production regulated by juvenile hormone Barth and Lester (1973) and Schal and Smith (1990) reviewed the early literature on hormone involvement in pheromone production in cockroaches. With the exception of Nauphoeta (detailed above), no studies on the regulation of volatile pheromone production are available that use analytical or biochemical approaches. The most detailed studies, with B. germanica and S. longipalpa, have employed behavioral and EAG responses of males to estimate the relative amount of pheromone in females or their pheromone glands. In both species, virgin females initiate pheromone production 4 days after the imaginal molt, in relation to increasing titers of JH III (Smith and Schal, 1990a; Liang and Schal, 1993). Ablation of the corpus allatum (CA) of newly emerged adult females prevents pheromone production in both species, and pheromone production is restored after reimplantation of active CA or by treatment with JH III or JH analogs. Interestingly, although growth of the vitellogenic oocytes is controlled by and highly correlated with JH III titers, direct or even intermediary involvement of the ovaries in regulating pheromone production and calling behavior in both species was excluded by ovariectomies (Smith and Schal, 1990a). It is not known whether JH exerts its pheromonotropic effects directly on mature secretory cells of

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the pheromone gland or if it acts indirectly by stimulating the synthesis and/or release of pheromonotropic neuropeptides. Although cockroach brain extracts induce PBAN-like pheromonotropic activity in moth pheromone glands (Raina et al., 1989), they fail to do so in allatectomized Nauphoeta males (Sreng et al., 1999) or Supella females (Schal, unpublished data). Moreover, lack of pheromone production in mated females that periodically produce large amounts of JH III suggests that JH plays a ‘‘permissive’’ role (Smith and Schal, 1990b; Schal et al., 1996). That is, its presence is required for pheromone to be produced, but even when the JH III titer is high pheromone production can be suppressed by neural or humoral pheromonostatic factors. Blattella germanica has served as a useful model for delineating endocrine regulation of nonvolatile cuticular pheromones, namely (3S,11S)-dimethylnonacosan-2-one. Both the amount of pheromone on the cuticular surface and in vivo incorporation of radiolabel from [14C1]propionate into the sex pheromone coincide with active stages of vitellogenesis, suggesting the involvement of JH (Schal et al., 1990a, 1994; Sevala et al., 1999). Indeed, females treated so as to reduce their JH III titer (allatectomy, antiallatal drugs such as precocene, starvation, or implantation of an artificial egg case into the genital vestibulum, which inhibits JH biosynthesis) produce less pheromone (Schal et al., 1990a, 1994; Chase et al., 1992). Furthermore, pheromone production is greatly stimulated by treatments with JH III or with JH analogs. Because only the hydroxylation of 3,11-dimethylnonacosane to 3,11-dimethylnonacosan-2-ol is regulated in a sex-specific manner, it appears that this step is under JH III control (Figure 11) (Chase et al., 1992). Normally, adult male cockroaches have a much lower titer of JH III in the hemolymph (Piulachs et al., 1992; review: Wyatt and Davey, 1996). Because males also produce 3,11-dimethylnonacosane, metabolism of the alkane to contact pheromone may be contingent upon high JH titers in the adult female. In response to exposure to the JH analog hydroprene, female pheromone increased sixfold in treated males (Schal, 1988), showing some capacity to express the putative female-specific polysubstrate monooxigenase. The parallels are striking with estrogen induction of vitellogenin synthesis in the male liver of oviparous vertebrates, JH induction of vitellogenin synthesis in male cockroaches (Mundall et al., 1983), and ecdysteroid induction of female pheromone production in houseflies (see Section 3.14.5.3.2) (Blomquist et al., 1984b, 1987).

Contact pheromone production in B. germanica is also regulated through the regulated production of its precursor, 3,11-dimethylnonacosane. Biosynthesis of this alkane drops dramatically when food intake declines at the end of each vitellogenic phase (Schal et al., 1994, 1996), suggesting that hydrocarbon biosynthesis is linked to food intake, as in nymphs (Young et al., 1999), and not directly to either the ecdysteroid or JH titers. Dietary intake also stimulates the production of JH III (Schal et al., 1993; Osorio et al., 1998), which in turn stimulates the conversion of the alkane to contact pheromone. In allatectomized females large amounts of hydrocarbons accumulate in the hemolymph because food intake is not suppressed (i.e., no gestation) and hydrocarbons are not provisioned into oocytes (i.e., no vitellogenesis) (Schal et al., 1994; Fan et al., 2002). As hydrocarbons accumulate in the hemolymph, the amount of cuticular pheromone also increases, suggesting that excess 3,11-dimethylnonacosane is metabolized to pheromone. These patterns suggest that under normal conditions, feeding in adult females is modulated in a stagespecific manner, regulating the amount of 3,11-dimethylnonacosane that is available for JHmediated metabolism of 3,11-dimethylnonacosane to 3,11-dimethylnonacosan-2-one. 3.14.5.5.3. Transport and emission of pheromones Little is known of the cellular processes that deliver volatile pheromones from secretory cells to the cuticular surface, even in the intensively researched Lepidoptera. In cockroaches, electron microscopy studies often show accumulation of secretion in the end-apparatus, ducts, and around the cuticular pores of class-3 exocrine glands in both males and females. Recent studies in L. maderae have identified and sequenced an epicuticular protein, Lma-p54, that is expressed specifically in the tergites and sternites of adult males and females, but not in nymphs (Cornette et al., 2002). The sequence of this protein is closely related to aspartic proteases, but because it appears to be enzymatically inactive the authors speculate that it serves as a ligandbinding protein. Cornette et al. (2002) further hypothesize that Lma-p54, alone or together with a ligand, serves in sexual recognition. Other ligandbinding proteins, namely the lipocalins Lma-p22 and Lma-p18, have been isolated only from male tergal secretions of L. maderae (Cornette et al., 2001). These exciting findings suggest that carrier proteins might be involved in the transport of volatile pheromones to the cuticle, but functional studies will be needed to verify this hypothesis.

Biochemistry and Molecular Biology of Pheromone Production

Attractant sex pheromones are usually emitted while the female or male cockroach performs a species-specific calling behavior (Gemeno and Schal, 2004; Gemeno et al., 2003). As in most lepidopterans, JH III regulates calling behavior in both species B. germanica and S. longipalpa. In both species, transection of the nerves connecting the CA to the brain, an operation that significantly accelerates the rate of JH III biosynthesis by the CA (Schal et al., 1993), also hastens the age when calling first occurs (Smith and Schal, 1990a; Liang and Schal, 1994). In B. germanica, the central role of JH has been confirmed by ablation of the CA and with rescue experiments with a JH analog (Liang and Schal, 1994). In this species, JH is also required for females to become sexually receptive and accept courting males (Schal and Chiang, 1995). In B. germanica, the transfer of contact pheromones to the epicuticular surface is mediated by lipophorin, a high-density hemolymph lipoprotein (reviews: Schal et al., 1998b, 2003). As in M. domestica (see Section 3.14.5.4.4), newly biosynthesized pheromone appears first in the hemolymph before it is noted on the epicuticle (Gu et al., 1995). That hemolymph is required to transport the pheromone to the cuticular surface was demonstrated by severing the veins that enter the forewings. Subsequently, the amount of hydrocarbons and pheromone that appeared on the wings was significantly lower than on the intact forewings of the same insects. In the cockroaches P. americana and B. germanica, virtually all newly synthesized hydrocarbons that enter the hemolymph are bound to lipophorin (Chino, 1985; Gu et al., 1995). Moreover, in both species newly synthesized hydrocarbons can only be transferred from the integument to an incubation medium if lipophorin is present, and other hemolymph lipoproteins, such as vitellogenin, cannot mediate this transfer (Katase and Chino, 1982, 1984; Fan et al., 2002). The mechanisms by which hydrocarbons and pheromones are taken up by lipophorin are poorly understood. Takeuchi and Chino (1993) demonstrated clearly in the American cockroach that a very-high-density lipid transfer particle (LTP) catalyzes the transfer of hydrocarbons between lipophorin particles. However, in vitro experiments with purified lipophorin of P. americana and B. germanica have shown that lipophorin accepts hydrocarbons from oenocytes, apparently without the involvement of LTP (Katase and Chino, 1982, 1984; Fan et al., 2002). It remains to be determined whether this is because sufficient LTP remains bound to dissected tissues, if LTP is produced by the dissected tissues, or whether it plays no

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significant role in hydrocarbon and pheromone uptake by lipophorin. Interestingly, while the uptake of hydrocarbons and pheromones by lipophorin in vitro appears to lack molecular specificity (Katase and Chino, 1984; Fan and Schal, unpublished data), their delivery to pheromone gland cells is highly specific (Schal et al., 1998a, 1998b). Uptake of lipophorin and its ligands might involve receptor-mediated endocytosis, as demonstrated in mosquito oocytes (Cheon et al., 2001; see Chapter 3.9). This might explain why some 3,11-dimethylnonacosan-2-one has been isolated from mature ovaries of the German cockroach (Gu et al., 1995) and (Z)-9-tricosene is found in housefly ovaries (Schal et al., 2001). However, an endocytic process would fail to discriminate various ligands, suggesting that alternative mechanisms need to be investigated.

3.14.6. Concluding Remarks and Future Directions Our increased understanding of the biochemistry and regulation of pheromone production over the last two decades has been most impressive. A 1983 review of the biochemistry and endocrine regulation of insect pheromone production (Blomquist and Dillwith, 1983) was 16 pages long. It was limited to early work on pheromone biosynthesis in moths and the housefly, and recognized that JH and ecdysteroids may play a role in the regulation of pheromone production. Since that time the discovery of PBAN and its role in the regulation of lepidopteran pheromone production have been elucidated, along with determining which enzymes are affected by JH and ecdysteroids to regulate pheromone production in model cockroaches/beetles and flies, respectively. The work in Lepidoptera has moved from simply demonstrating that pheromone components were synthesized de novo to the molecular characterization of the unique D11-desaturase and other desaturases that are involved in many female moths and their interplay with specific chain shortening steps. While it is still true that in no system do we have a complete understanding of both the biochemical pathways and their endocrine regulation, we do have a much better understanding of how pheromones are made and in some systems are developing an understanding of their regulation at the molecular level. The continued application of the powerful tools of molecular biology along with studies using genomics and proteomics will only increase the rate at which we increase our understanding of pheromone production. Ultimately, just as behavioral chemicals themselves have been extended into pest

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control, research on pheromone production will be directed toward practical applications in insect control.

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Schal, C., Sevala, V., Capurro, M.d.L., et al., 2001. Tissue distribution and lipophorin transport of hydrocarbon and sex pheromones in the house fly, Musca domestica. J. Insect Sci. 1, 12 Available online:. insectscience.org/1.12. Schal, C., Sevala, V., Carde´, R.T., 1998a. Novel and highly specific transport of a volatile sex pheromone by hemolymph lipophorin in moths. Naturwiss. 85, 339–342. Schal, C., Sevala, V.L., Young, H.P., Bachmann, J.A.S., 1998b. Synthesis and transport of hydrocarbons: cuticle and ovary as target tissues. Am. Zool. 38, 382–393. Schal, C., Smith, A.F., 1990. Neuroendocrine regulation of pheromone production in cockoraches. In: Huber, I., Masler, E.P., Rao, B.R. (Eds.), Cockroaches as Models for Neurobiology: Applications in Biomedical Research. CRC Press, Boca Raton, FL, pp. 179–200. Schlyter, F., Birgersson, G.S., 1999. Forest beetles. In: Hardie, J., Minks, A.K. (Eds.), Pheromones of NonLepidopteran Insects Associated with Agricultural Plants. CAB International, Wallingford, pp. 113–148. Schoofs, L., Holman, G.M., Hayes, T.K., Nachman, R.J., DeLoof, A., 1991. Isolation, primary structure and synthesis of locustapyrokinin: a myotropic peptide of Locusta migratoria. Gen. Comp. Endocrinol. 81, 97–104. Schoofs, L., Holman, G.M., Hayes, T.K., Nachman, R.J., Kochansky, J.P., et al., 1992. Isolation, identification and synthesis of locustamyotropin III and IV, two additional neuropeptides of Locusta migratoria: members of the locustamyotropin peptide family. Insect Biochem. Mol. Biol. 22, 447–452. Schoofs, L., Holman, G.M., Hayes, T.K., Nachman, R.J., Loof, A.D., 1990a. Isolation, identification and synthesis of locustamyotropin II, an additional neuropeptide of Locusta migratoria: member of the cephalomyotropic peptide family. Insect Biochem. 20, 479–484. Schoofs, L., Holman, G.M., Hayes, T.K., et al., 1990b. Isolation, identification and synthesis of locustamyotropin (Lom-MT), a novel biologically active insect neuropeptide. Peptides 11, 427–433. Schoofs, L., Holman, G.M., Nachman, R., Proost, P., Vandamme, J., et al., 1993. Isolation, identification and synthesis of locustapyrokinin-II from Locusta migratoria, another member of the FXPRLamide peptide family. Comp. Biochem. Physiol. C 106, 103–109. Sevala, V., Shu, S., Ramaswamy, S.B., Schal, C., 1999. Lipophorin of female Blattella germanica (L.): characterization and relation to hemolymph titers of juvenile hormone and hydrocarbons. J. Insect Physiol. 45, 431–441. Seybold, S.J., Bohlmann, J., Raffa, K.F., 2000. Biosynthesis of coniferophagous bark beetle pheromones and conifer isoprenoids: evolutionary perspective and synthesis. Can. Entomol. 132, 697–753. Seybold, S.J., Ohtsuka, T., Wood, D.L., Kubo, I., 1995a. Enantiomeric composition of ipsdienol: a chemotaxonomic character for North American populations of Ips spp. in the pini subgeneric group (Coleoptera: Scolytidae). J. Chem. Ecol. 21, 995–1016.

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Seybold, S.J., Quilici, D.R., Tillman, J.A., Vanderwel, D., Wood, D.L., et al., 1995b. De novo biosynthesis of the aggregation pheromone components ipsenol and ipsdienol by the pine bark beetles Ips paraconfusus Lanier and Ips pini (Say) (Coleoptera: Scolytidae). Proc. Natl Acad. Sci. USA 92, 8393–8397. Seybold, S.J., Tittiger, C., 2003. Biochemistry and molecular biology of de novo isoprenoid pheromone production. 1. The Scolytidae. Annu. Rev. Entomol. 48, 425–453. Seybold, S.J., Vanderwel, D., 2003. Biosynthesis and endocrine regulation of pheromone production in the Coleoptera. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Pheromone Biochemistry and Molecular Biochemistry. Elsevier, San Diego, CA, pp. 137–200. Shorey, H.H., 1974. Environmental and physiological control of insext sex pheromone behavior. In: Birch, M.C. (Ed.), Pheromones. Elsevier, New York, pp. 62–80. Silverstein, R.M., Rodin, J.O., Wood, D.L., 1966. Sex attractants in frass produced by male Ips confusus in ponderosa pine. Science 154, 509–510. Sirugue, D., Bonnard, O., Le Quere, J.L., Farine, J.-P., Brossut, R., 1992. 2-Methylthiazolidine and 4-ethylguaiacol, male sex pheromone components of the cockroach Nauphoeta cinerea (Dictyoptera, Blaberidae): a reinvestigation. J. Chem. Ecol. 18, 2261–2276. Smith, A.F., Schal, C., 1990a. Corpus allatum control of sex pheromone production and calling in the female brown-banded cockroach, Supella longipalpa (F.) (Dictyoptera: Blattellidae). J. Insect Physiol. 36, 251–257. Smith, A.F., Schal, C., 1990b. The physiological basis for the termination of pheromone-releasing behaviour in the female brown-banded cockroach, Supella longipalpa (F.) (Dictyoptera: Blattellidae). J. Insect Physiol. 36, 369–373. Smith, V.L., Doyle, K.E., Maune, J.F., Munjaal, R.P., Beckingham, K., 1987. Structure and sequence of the Drosophila melanogaster calmodulin gene. J. Mol. Biol. 196, 471–485. Soroker, V., Rafaeli, A., 1989. In vitro hormonal stimulation of [14C]acetate incorporation by Heliothis armigera pheromone glands. Insect Biochem. 19, 1–5. Sreng, L., 1990. Seducin, male sex pheromone of the cockroach Nauphoeta cinerea: isolation, identification, and bioassay. J. Chem. Ecol. 16, 2899–2912. Sreng, L., 1998. Apostosis-inducing brain factors in maturation of an insect sex pheromone gland during differentiation. Differentiation 63, 53–58. Sreng, I., Glover, T., Roelofs, W., 1989. Canalization of the redbanded leafroller moth sex pheromone blend. Arch. Insect Biochem. Physiol. 10, 73–82. Sreng, L., Leoncini, I., Clement, J.L., 1999. Regulation of sex pheromone production in the male Nauphoeta cinerea cockroach: role of brain extracts, corpora allata (CA), and juvenile hormone (JH). Arch. Insect Biochem. Physiol. 40, 165–172. Sreng, L., Quennedey, A., 1976. Role of a temporary ciliary structure in the morphogenesis of insect glands. An electron microscope study of the tergal glands of

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male Blattella germanica L. (Dictyoptera, Blattellidae). J. Ultrastruct. Res. 56, 78–95. Stanley-Samuelson, D.W., Jurenka, R.A., Cripps, C., Blomquist, G.J., deRenobales, M., 1988. Fatty acids in insects: composition, metabolism and biological significance. Arch. Insect Biochem. Physiol. 9, 1–33. Subchev, M., Jurenka, R.A., 2001. Identification of the pheromone in the hemolymph and cuticular hydrocarbons from the moth Scoliopteryx libatrix L. (Lepidoptera: Noctuidae). Arch. Insect Biochem. Physiol. 47, 35–43. Tada, S., Leal, W.S., 1997. Localization and morphology of sex pheromone glands in scarab beetles. J. Chem. Ecol. 23, 903–915. Takeuchi, N., Chino, H., 1993. Lipid transfer particle in the hemolymph of the American cockroach: evidence for its capacity to transfer hydrocarbons between lipophorin particles. J. Lipid Res. 34, 543–551. Tamaki, Y., 1985. Sex pheromones. In: Kerkut, G.A., Gilbert, L.I. (Eds.), Comprehensive Insect Physiology, Biochemistry and Pharmacology, vol. 9. Pergamon Press, Oxford, pp. 145–191. Tang, J.D., Charlton, R.E., Carde´ , R.T., Yin, C.-M., 1987. Effect of allatectomy and ventral nerve cord transection on calling, pheromone emission and pheromone production in Lymantria dispar. J. Insect Physiol. 33, 469–476. Tang, J.D., Charlton, R.E., Jurenka, R.A., Wolf, W.A., Phelan, P.L., et al., 1989. Regulation of pheromone biosynthesis by a brain hormone in two moth species. Proc. Natl Acad. Sci. USA 86, 1806–1810. Teal, P.E.A., Tumlinson, J.H., 1987. The role of alcohols in pheromone biosynthesis by two noctuid moths that use acetate pheromone components. Arch. Insect Biochem. Physiol. 4, 261–269. Teal, P.E.A., Tumlinson, J.H., 1988. Properties of cuticular oxidases used for sex pheromone biosynthesis by Heliothis zea. J. Chem. Ecol. 14, 2131–2145. Thompson, A.C., Mitlin, N., 1979. Biosynthesis of the sex pheromone of the male boll weevil from monoterpene precursors. Insect Biochem. 9, 293–294. Thyagaraja, B.S., Raina, A.K., 1994. Regulation of pheromone production in the gypsy moth, Lymantria dispar, and development of an in vitro bioassay. J. Insect Physiol. 40, 969–974. Tillman, J.A., Holbrook, G.L., Dallara, P., Schal, C., Wood, D.L., et al., 1998. Endocrine regulation of de novo aggregation pheromone biosynthesis in the pine engraver, Ips pini (Say) (Coleoptera: Scolytidae). Insect Biochem. Mol. Biol. 28, 705–715. Tillman, J.A., Seybold, S.J., Jurenka, R.A., Blomquist, G.J., 1999. Insect pheromones: an overview of biosynthesis and endocrine regulation. Insect Biochem. Mol. Biol. 29, 481–514. Tillman-Wall, J.A., Vanderwel, D., Kuenzli, M.E., Reitz, R.C., Blomquist, G.J., 1992. Regulation of sex pheromone biosynthesis in the housefly, Musca domestica: relative contribution of the elongation and reductive step. Arch. Biochem. Biophys. 299, 92–99.

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Tittiger, C., 2003. Molecular biology of bank beetle pheromone production and endocrine regulation. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Biochemistry and Molecular Biology. Elsevier, San Diego, CA, pp. 201–230. Tittiger, C., Barkawi, L.S., Bengoa, C.S., Blomquist, G.J., Seybold, S.J., 2003. Structure and juvenile hormonemediated regulation of the HMG-CoA reductase gene from the Jeffrey pine beetle, Dendroctonus jeffreyi. Mol. Cell. Endocrinol. 199, 11–21. Tittiger, C., Blomquist, G.J., Ivarsson, P., Borgeson, C.E., Seybold, S.J., 1999. Juvenile hormone regulation of HMG-R gene expression in the bark beetle Ips paraconfusus (Coleoptera: Scolytidae): implications for male aggregation pheromone biosynthesis. Cell. Mol. Life Sci. 55, 121–127. Tittiger, C., O’Keeffe, C., Bengoa, C.S., et al., 2000. Isolation and endocrine regulation of HMG-CoA synthase cDNA from the male Jeffrey pine beetle, Dendroctonus jeffreyi. Insect Biochem. Mol. Biol. 30, 1203–1211. Tokro, P.G., Brossut, R., Sreng, L., 1993. Studies on the sex pheromone of female Blattella germanica L. Insect Sci. Applic. 14, 115–126. Torfs, P., Nieto, J., Cerstiaens, A., Boon, D., Baggerman, G., et al., 2001. Pyrokinin neuropeptides in a crustacean: isolation and identification in the white shrimp Penaeus vannamei. Eur. J. Biochem. 268, 149–154. Tumlinson, J.H., Fang, N., Teal, P.E.A., 1997. The effect of PBAN on conversion of fatty acyls to pheromone aldehydes in female. In: Carde´ , R.T., Minks, A.K. (Eds.), Insect Pheromone Research: New Directions. Chapman and Hall, New York, pp. 54–55. Vanderwel, D., 1994. Factors affecting pheromone production in beetles. Arch. Insect Biochem. Physiol. 25, 347–362. Vanderwel, D., Oehlschlager, A.C., 1987. Biosynthesis of pheromones and endocrine regulation of pheromone production in Coleoptera. In: Prestwich, G.D., Blomquist, G.J. (Eds.), Pheromone Biochemistry. Academic Press, Orlando, FL, pp. 175–215. Vaz, A.H., Jurenka, R.A., Blomquist, G.J., Reitz, R.C., 1988. Tissue and chain length specificity of the fatty acyl-CoA elongation system in the American cockroach. Arch. Biochem. Biophys. 267, 551–557. Veelaert, D., Schoofs, L., Verhaert, P., De Loof, A., 1997. Identification of two novel peptides from the central nervous system of the desert locust, Schistocerca gregaria. Biochem. Biophys. Res. Commun. 241, 530–534. Wakayama, E.J., Dillwith, J.W., Howard, R.W., Blomquist, G.J., 1984. Vitamin B12 levels in selected insects. Insect Biochem. 14, 175–179. Wei, W., Miyamoto, T., Endo, M., et al., 2003. Polyunsaturated hydrocarbons in the hemolymph: biosynthetic precursors of epoxy pheromones of geometrid and arctiid moths. Insect Biochem. and Mol. Biol. 33, 397–405.

Wicker, C., Jallon, J.M., 1995a. Influence of ovary and ecdysteroids on pheromone biosynthesis in Drosophila melanogaster (Diptera, Drosophilidae). Eur. J. Entomol. 92, 197–202. Wicker, C., Jallon, J.-M., 1995b. Hormonal control of sex pheromone biosynthesis in Drosophila melanogaster. J. Insect Physiol. 41, 65–70. Wicker-Thomas, C., Henriet, C., Dallerac, R., 1997. Partial characterization of a fatty acid desaturase gene in Drosophila melanogaster. Insect Biochem. Mol. Biol. 11, 963–972. Wiygul, G., Dickens, J.C., Smith, J.W., 1990. Effect of juvenile hormone and b-bisabolol on pheromone production in fat bodies of male boll weevils, Anthonomus grandis Boheman (Coleoptera; Curculionidae). Comp. Biochem. Physiol. B 95, 489–491. Wolf, W.A., Roelofs, W.L., 1983. A chain-shortening reaction in orange tortrix moth sex pheromone biosynthesis. Insect Biochem. 13, 375–379. Wolf, W.A., Roelofs, W.L., 1986. Properties of the D11desaturase enzyme used in cabbage looper moth sex pheromone biosynthesis. Arch. Insect Biochem. Physiol. 3, 45–52. Wolf, W.A., Roelofs, W.L., 1987. Reinvestigation confirms action of D11-desaturase in spruce budworm moth sex pheromone biosynthesis. J. Chem. Ecol. 13, 1019–1027. Wood, D.L., 1982. The role of pheromones, kairomones, and allomones in the host selection and colonization behavior of bark beetles Coleoptera. Ann. Rev. Entomol. 27, 411–446. Wu, W.Q., Zhu, J.W., Millar, J., Lo¨ fstedt, C., 1998. A comparative study of sex pheromone biosynthesis in two strains of the turnip moth, Agrotis segetum, producing different ratios of sex pheromone components. Insect Biochem. Mol. Biol. 28, 895–900. Wyatt, G.R., Davey, K.G., 1996. Cellular and molecular actions of juvenile hormone. 2. Roles of juvenile hormone in adult insects. Adv. Insect Physiol. 26, 1–155. Yang, H.-T., Chow, Y.-S., Peng, W.-K., Hsu, E.-L., 1998. Evidence for the site of female sex pheromone production in Periplaneta americana. J. Chem. Ecol. 24, 1831–1843. Yoder, J.A., Denlinger, D.L., Dennis, M.W., Kolattukudy, P.E., 1992. Enhancement of diapausing flesh fly puparia with additional hydrocarbons and evidence for alkane biosynthesis by a decarbonylation mechanism. Insect Biochem. Mol. Biol. 22, 237–243. Yoshiga, T., Yokoyama, N., Imai, N., Ohnishi, A., Moto, K., et al., 2002. cDNA cloning of calcineurin heterosubunits from the pheromone gland of the silkmoth, Bombyx mori. Insect Biochem. Mol. Biol. 32, 477–486. Young, H.P., Bachmann, J.A.S., Schal, C., 1999. Food intake in Blattella germanica (L.) nymphs affects hydrocarbon synthesis and its allocation in adults between epicuticle and reproduction. Arch. Insect Biochem. Physiol. 41, 214–224.

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Zdarek, J., Nachman, R.J., Hayes, T.K., 1998. Structure– activity relationships of insect neuropeptides of the pyrokinin/PBAN family and their selective action on pupariation in fleshfly (Neobelleria bullata) larvae (Diptera, Sarcophagidae). Eur. J. Entomol. 95, 9–16. Zhang, T., Zhang, L., Xu, W., Shen, J., 2001. Cloning and characterization of the cDNA of diapause hormone–pheromone biosynthesis activating neuropeptide of Helicoverpa armigera. GenBank Direct Submission. Zhao, C., Lo¨ fstedt, C., Wang, X., 1990. Sex pheromone biosynthesis in the Asian corn borer Ostrinia furnicalis 2. Biosynthesis of (E) and (Z)-12-tetradecenyl acetate involves D14 desaturation. Arch. Biochem. Physiol. 15, 57–65.

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Zhu, J., Millar, J., Lo¨ fstedt, C., 1995. Hormonal regulation of sex pheromone biosynthesis in the turnip moth, Agrotis segetum. Arch. Biochem. Physiol. 30, 41–59. Zhu, J., Zhao, C.H, Lu, F., Bengtsson, M., Lo¨ fstedt, C., 1996. Reductase specificity and the ratio regulation of E/Z isomers in pheromone biosynthesis of the European corn borer, Ostrinia nubilalis (Lepidoptera: Pyralidae). Insect Biochem. Mol. Biol. 26, 171–176. Relevant Website http://www.nysaes.cornell.edu – The Phenolist, a listing of lepidopteran phenomones compiled at Cornell University.

3.15

Molecular Basis of Pheromone Detection in Insects

R G Vogt, University of South Carolina, Columbia, SC, USA ß 2005, Elsevier BV. All Rights Reserved.

3.15.1. Introduction 3.15.2. Molecular Basis of Insect Chemodetection: General Schemes 3.15.3. Transduction Events 3.15.3.1. Receptors 3.15.3.2. Second Messenger Pathways 3.15.4. Perireception Events 3.15.4.1. Odor Transport by Odorant Binding Proteins 3.15.4.2. Odor Transport by OS-Ds, SAPs, or CSPs? 3.15.4.3. Pheromone and Odor Degradation by Odor Degrading Enzymes 3.15.4.4. Sensory Neuron Membrane Proteins 3.15.4.5. Perireceptor Models 3.15.5. Conclusions 3.15.5.1. Does the Detection of Pheromones Differ from the Detection of Other Odor Molecules? 3.15.5.2. Are OBPs Necessary for OR Activation? 3.15.5.3. Regulation of Pheromone Detection by Hormones, and the Regulation of Hormones by Pheromones

3.15.1. Introduction The detection of environmental chemicals is surely one of the oldest senses, critical to bacteria, singlecelled eukaryotes, plants, fungi, and animals. Chemical signatures carry enormously diverse information about the external world. Pheromones represent one class of environmental signals and are stringently defined as chemicals produced by individuals and detected by other members of the same species (Karlson and Lu¨scher, 1959). Pheromones have been categorized as primers, stimulating or modulating an immediate behavioral response, and releasers, stimulating or modulating a longer-lasting physiological state (Birch, 1974; Wyatt, 2003). Pheromone detection is viewed as among the most specialized form of chemodetection, and the mechanisms underlying the detection and processing of pheromone signals have been viewed not merely as a comprising a special case of chemodetection, but often as a distinct process. However, chemicals produced outside the species may have effects as profound as pheromones on an individual, either short term or long term. Furthermore, the underlying mechanisms for detecting and processing pheromone and nonpheromone chemical signals are turning out to be quite similar (e.g., Christensen and Hildebrand, 2002). Certain chemical signals, including pheromones but also karimones (signals between species) and plant-derived oviposition cues,

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are components of processes that have special roles in the life of an organism, and as such, evolutionary selection has acted to elaborate preexisting detection/response pathways in ways appropriate for the processing of those chemical signals. So, while pheromone detection pathways are clearly under different kinds of selective pressure than say food-odor detection pathways, the specializations seen in pheromone pathways are most likely modifications of an overall common process. Nevertheless, pheromones are clearly unique among all chemical signals, in that selection has acted on different members of the same species to coordinate both the production and detection of these signals. And yet, remarkably much is still unexplored in the area of the physiological/hormonal regulation of these two distinct pheromonal processes. How are pheromones detected? Insects have two major chemosensory systems, olfaction (smell) and gustation (taste) (Stocker, 1994). Pheromones might be detected by either system, though virtually all that is known about the mechanisms underlying pheromone detection involves olfaction. Olfaction might be defined as the detection of volatile chemostimulants, but a more appropriate definition relates to the unique neuroanatomy of olfaction. In insects, odors are detected by olfactory sensilla which come in a variety of shapes, including long and short hair-like structures and plate-like

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Figure 1 Frontal view of an adult insect brain detailing olfactory pathways through the brain. Left: Dendrites and cell bodies of olfactory sensory neurons (OSNs) are located in the antennas (ANT) and palps within sensilla; axons project, via the antennal nerve (AN), to spherical glomerulae in the antennal lobe (AL) where they synapse with local interneurons (LINs) and projection neurons (PNs). OSNs expressing the same OR project to the same glomerulus (e.g., neurons A) while OSNs expressing different receptors project to different glomerulae (e.g., neurons B, C) (Hansson et al., 1992, 2003; Anton and Hansson, 1995; Joerges et al., 1997; Galizia et al., 1998, 1999, 2000; Gao et al., 2000; Vosshall et al., 2000; Galizia and Menzel, 2001; Carlsson et al., 2002; Sadek et al., 2002; Marin et al., 2002; Carlsson and Hansson, 2003a, 2003b). Right: The AL contains multiple types of glomerulae, including specialized glomerulae such as the macroglomerular complex (MGC) (male specific in Lepidoptera) which comprises several glomerular units each receiving input from OSNs responding to specific components of a sex-pheromone blend (e.g., Christensen and Hildebrand, 2002; Hansson, 2002). LINs and PNs project from cell bodies located in one of several cell clusters (CC) located just outside the glomerular mass. LINs are multiglomerular, innervating all glomerulae, only the non-MGC glomerulae, or only the MGC (Matsumoto and Hildebrand, 1981; Hansson et al., 1994). PNs are uniglomerular and projecting outward to the lateral protocerebrum (LPC) via one of two tracks. PNs of the inner antenno-cerebral tract (i-ACT) make synaptic connections with neurons of the mushroom bodies (MB) as well as in the LPC, while PNs of the outer antenno-cerebral tract (o-ACT) only make synaptic connections in the LPC (Homberg et al., 1988; Kanzaki et al., 1989, 2003; Hansson et al., 1991; Sun et al., 1997; McGuire et al., 2001). Individual PNs innervate spatially restricted regions of the LPC, suggesting that there is a spatial conservation between glomerulus and LPC domain (Wong et al., 2002).

structures, and which may have single or double cuticular walls; but all are multiporous, having many small holes penetrating the cuticle to provide odor molecules access to chemosensory neurons within (Steinbrecht, 1997, 1999; Boeckh et al., 1965; see also Schneider, 1969; Kaissling, 1971). Adult olfactory neurons project to the olfactory lobes in the brain (Hildebrand, 1996; Hansson et al., 2003) (Figure 1); larval olfactory neurons project to a location in the brain homologous to the adult olfactory lobe (Kent and Hildebrand, 1987; Python and Stocker, 2002). The dendrites of olfactory neurons are ensheathed within cuticular hairs, comprising olfactory sensilla which are restricted to the head (primarily antennas and maxillary palps) (Steinbrecht, 1997, 1999). While these olfactory sensilla certainly detect volatile chemicals

for terrestrial insects, there are no data to say that homologous structures, as opposed to contact or gustatory sensilla, do not detect water-soluble molecules for aquatic insects. Nevertheless, it is the axonal projection pathway of the chemsensory neurons to the olfactory lobe that most consistently distinguishes the olfactory from the gustatory system. Gustation might be defined as the detection of nonvolatile chemostimulants, but again, an anatomically based definition may better serve. Gustatory sensilla have a single pore at the tip (Steinbrecht, 1984); their frequent description as contact chemoreceptors confers the idea that these sensilla must make physical contact with a substrate permitting a chemostimulant to flow into the pore and to the sensory neurons within. Gustatory sensilla typically contain several chemosensory neurons tuned to

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various modalities which may include nutrients, feeding stimulants or deterrents, water and/or salts, and may also include a mechanosensitive neuron (Hanson and Dethier, 1973; Steinbrecht, 1984; Kent and Hildebrand, 1987; Singh, 1997; Shanbhag et al., 2001b). Neurons of these sensilla located on the head project to the subesophageal ganglia (Kent and Hildebrand, 1987; Dunipace et al., 2001; Scott et al., 2001). Gustatory sensilla are also situated elsewhere on the insect body, including legs, wings, and genitalia, and chemosensory axons from these locations project to the appropriate ganglion or ganglionic region serving that segment (Stocker, 1994). In general, neurons of gustatory sensilla do not project to the olfactory lobe, again providing a distinction between these two systems. However, a very small number of the gustatory receptor genes recently identified in Drosophila melanogaster have been shown to express in neurons that do project to the olfactory lobe (Scott et al., 2001), emphasizing, perhaps, that in biology, no human-made distinctions should be set in stone. There seems no reason that insect pheromones should be detected exclusively by olfactory rather than gustatory sensilla, although the majority of those pheromones that have been characterized are processed by the olfactory system. Nevertheless, pheromones run the gamut of volatility, including the relatively nonvolatile cuticular hydrocarbon pheromones of dipteran and various social insects (Blomquist et al., 1998; Blomquist, 2003) (see Chapter 3.14). Also, there are numerous examples of insect courtship that employ both pheromones of low volatility and physical contact, suggesting that physical contact may be required for pheromone transfer, e.g. orthopteroids (Tregenza and Wedell, 1997; Rivault et al., 1998) and social insects (review: Clement and Bagneres, 1998). Drosophila employs several types of pheromones. Several peptides are transferred with the spermatophore during mating which may enter the hemolymph and bind to olfactory axons to modulate olfactory response (Ottiger et al., 2000) (see Chapter 1.5). A volatile aggregation pheromone, cis-vaccenyl acetate, is deposited on eggs and detected by trichoid sensilla on the antennas (Bartelt et al., 1985; Clyne et al., 1997). Various cuticular hydrocarbons also influence courtship behavior, including female specific cis,cis-7,11-heptacosadiene which alters male wing-beat frequency, and others which inhibit male–male interaction (Ferveur et al., 1996, 1997; Ferveur, 1997; Savarit et al., 1999). The Drosophila cuticular hydrocarbon pheromones have been assumed to be detected by contact

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gustatory sensilla (Greenspan and Ferveur, 2000). In particular, Drosophila prothoracic legs show a sexual dimorphism in the number of gustatory sensilla, with males having the greater number (Nayak and Singh, 1983; Meunier et al., 2000). This sexual asymmetry in sensilla number has led to suggestions that these sensilla may contain neurons that respond to the cuticular hydrocarbon pheromones (Robertson, 1983; Venard et al., 1989; Stocker, 1994; de Bruyne, 2003). These sensilla have yet to be evaluated for the physiological response to cuticular hydrocarbons. However, they have been shown to respond to sugars, salts, and amino acids (the expected stimulants of the respective chemosensory neurons) and furthermore to show a corresponding sexual asymmetry in their response to these nonpheromonal stimulants (Meunier et al., 2000), suggesting some sex-specific but nonpheromonal role for these gustatory sensilla. It may yet be demonstrated that Drosophila gustatory sensilla detect pheromones, but so far only olfactory sensilla of the antenna have been shown to participate in courtship behavior (Stocker and Gendre, 1989) or pheromone detection (Bartelt et al., 1985; Clyne et al., 1997). (Note: since this chapter was submitted, new data have been presented suggesting Drosophila leg gustatory sensilla are involved in pheromone detection, mediated by gustatory receptor gene Gr68a; see Bray and Amrein, 2003; Matsunami and Amrein, 2003.) It remains less than definitive that any insect pheromones are in fact detected by gustatory sensilla. However, gustatory sensilla have been shown to function as pheromone detectors in noninsect arthropods such as ticks (Sonenshine, 1985; de Bruyne and Guerin, 1998). The lack of information from insects may merely be a sampling bias, with the vast majority of molecular and physiological research effort being focused on relatively few types of olfactory based pheromones. This review covers the molecular biology of insect olfaction and to a lesser extent taste, with specific attention to those biochemical events, which led to the initial depolarization of chemosensory neurons. The discussion will be restricted to known gene products and the processes they mediate. In most cases, there are excellent examples of these processes working in the detection of pheromones. The exception to this is with odor receptors; as of early 2000s no pheromone receptor proteins have been reported, though this circumstance will likely change by the time this work is published, or very soon thereafter, as the genomes of the honeybee and several Lepidoptera become fully characterized.

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3.15.2. Molecular Basis of Insect Chemodetection: General Schemes Figure 2 summarizes our current general understanding of the biochemical events involved in the detection of pheromones and other odors by insect olfactory sensilla. Odor molecules enter the aqueous lumen of a sensillum via pores penetrating the cuticle (Steinbrecht, 1997; Steinbrecht and Stankiewicz, 1999). Once inside, odors are thought to bind to and be transported by soluble odorant binding proteins (OBPs) which deliver the odors to transmembrane olfactory receptor proteins (ORs). Odor molecules are subsequently degraded by a variety of enzymes that may be located within the sensillum lumen as well as within the cells lining the base of the sensillum. ORs so far identified are 7-transmembrane domain G-protein-coupled receptors (Vosshall and Keller, 2003); activation of these receptors is thought to generate a rapid increase in inositol triphosphate (IP3) which is thought to directly activate ion channels in the neuronal membrane. The Drosophila genome contains at least 61 OR genes and more than 50 OBP genes, providing the molecular basis for the detection of diverse odorants. For all but the ORs and arrestins, each of the biochemical events depicted in Figure 2 has examples in the processing of pheromone signals. The biochemical processes at work in gustatory sensilla are less well characterized. Gustatory sensilla generally contain multiple sensory neurons including several chemosensory neurons and often one mechanosensory neuron; the chemosensory neurons are tuned to different modalities allowing insects, as is the case with vertebrates, to distinguish sweet, sour, salty, and bitter substances (review: Singh, 1997). However, the range of chemicals that insects detect with these sensilla is likely quite broad, e.g., ecdysteroid chemodetection (Descoins and Marion-Poll, 1999). A family of at least 55 Gprotein coupled gustatory receptor (GR) genes has recently been identified in Drosophila (Clyne et al., 2000; Dunipace et al., 2001; Scott et al., 2001) and the malaria mosquito Anopheles gambiae (Hill et al., 2002). Of these, one (Gr5a) has been shown to interact with the sugar trehalose (Dahanukar et al., 2001; Ueno et al., 2001). An IP3 cascade, similar to that shown in Figure 2c, has also been described for the gustatory detection of sugars and amino acids (Koganezawa and Shimada, 2002). A member of the OBP family has also been reported to associate with certain gustatory sensilla of the blowfly Phormia regina, but participates in the detection of certain volatile compounds rather than those (e.g., sugars

and amino acids) typically associated with gustatory sensilla (Ozaki et al., 1995, 2003). The term ‘‘perireceptor events’’ was coined by Getchell et al. (1984) to describe processes in vertebrate olfaction that preceded the odor–OR interactions (see also Carr et al., 1990). The equivalent events in insect olfaction are depicted in Figure 2b, and are those involving OBPs, ODEs, and their interaction with ORs. Transductory events, then, are those depicted in Figure 2c, and involve ORs and the second messenger pathway(s) they regulate. While these two processes, perireception and transduction, clearly overlap at least at the level of the ORs, they remain useful distinctions. In particular, the transduction events that follow receptor activation appear relatively common for all odor–OR combinations. There may be more than one transduction pathway involved, as has been observed in the lobster (Panulirus argus) where both IP3 and cAMP pathways exist (Fadool and Ache, 1992; Michel and Ache, 1992; Boekhoff et al., 1994; Hatt and Ache, 1994). However, the immediate transduction elements (Figure 2c) do not show the genetic diversity seen for the perireceptor elements (Figure 2b). The OBPs, ODEs, and ORs are present in numbers and diverse sequences or types that suggest they are under strong evolutionary selective/diversification pressures characteristic of both the broad chemoreceptive needs and the divergent life histories of the species. The majority of the elements depicted in Figure 2 have been identified since the publication of the previous edition of this work. In the early 1980s there were few laboratories studying the biochemistry of odor detection in insects. In 1981 the first identification of a pheromone binding protein (PBP) and a sensillar esterase (SE) was reported; both appeared uniquely expressed in the male antennas of the silk moth Antheraea polyphemus and were present in the extracellular fluid of the sensillum lumen (Vogt and Riddiford, 1981). In 1985 a scheme was proposed in which pheromone molecules first bound to PBPs for transport to ORs and were subsequently rapidly degraded by SEs (Vogt et al., 1985). The perceived need for such transport arose because pheromone molecules are hydrophobic odors that should not readily enter the aqueous interior of the sensillum lumen. This new scheme was a dramatic departure from the prevailing view that pore tubules (anatomical structures of the cuticle wall) served as conduits for odors from air to neuron (reviews: Vogt, 1987; Steinbrecht, 1997). The subsequent identification of general odorant binding proteins (GOBP1 and GOBP2) suggested that this new scheme might be extended to other

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Figure 2 Schematic of an olfactory sensillum and a generalized biochemical pathway of odor reception. (a) Anatomy. An olfactory sensillum includes two or three neurons surrounded by three support cells; olfactory dendrites/cilia project up the fluid-filled lumen of a cuticular hair. The sensillum lumen is isolated from hemolymph by a cellular barrier. (Modified from Steinbrecht, R.A., 1969. Comparative morphology of olfactory receptor. In: Pfaffmann, C. (Ed.), Olfaction and Taste III. Rockefeller University Press, New York, pp. 3–21.) Olfactory sensilla come in a range of shapes, including plate-like and long and short hair-like structures, and olfactory neurons may have branched or unbranched dendrites; sensilla classes are in part defined by the shape of the cuticular hair and the branched nature of the neurons (Steinbrecht, 1997, 1999). There are little data on the functional differences between different sensilla classes of a given individual, although in Lepidoptera, attractant sex pheromones are detected by the long trichoid sensilla, illustrated here. (b) Perireceptor events. Hydrophobic odor molecules enter the aqueous sensillum lumen via pores penetrating the cuticular hair wall. Hydrophilic odorant binding proteins (OBPs) are proposed to bind and transport odors to receptor proteins located in the neuronal membranes. Odor degrading enzymes (ODEs) (pathway I) in the sensillum lumen are proposed to degrade these odor molecules. Cytoplasm of support cells contains xenobiotic inactivating enzymes (pathway IIa), such as glutathione-S-transferase (GST), which may also serve to inactivate odor molecules (pathway IIb). Interactions between OBPs and ORs and the function of sensory neuron membrane proteins (SNMPs) are unclear. (Modified from Rogers, M.E., Jani, M.K., Vogt, R.G., 1999. An olfactory specific glutathione S-transferase in the sphinx moth Manduca sexta. J. Exp. Biol. 202, 1625–1637.) (c) Transduction events. Odor–OR interaction stimulates an inositor triphosphate (IP3) second messenger cascade in which the a-subunit of a G-protein activates phospholipase C (PLC) to cleave the membrane lipid phosphotidyl inositol 4,5-bisphosphate (PIP2) to diacylglycerol (DAG) and IP3. IP3 is thought to bind directly to and activate transmembrane cation channels (IP3 receptors) (review: Krieger and Breer, 2003). Processes such as those mediated by phosphorylation (Schleicher et al., 1994) or arrestins (Merrill et al., 2002) may provide modulatory feedback on receptor–G-protein interactions. ‘‘?’’ pointing to Gbg, PIP2, and DAG are to imply yet uncharacterized but possible modulatory roles for these signals.

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olfactory modalities such as the detection of plant volatiles (Vogt and Lerner, 1989; Vogt et al., 1991a). The identification of an antennal specific but sex indifferent pheromone degrading aldehyde oxidase (AOX) in the moths Manduca sexta and A. polyphemus suggested that ODEs might also be a general feature of olfaction for both pheromones and plant volatiles (Rybczynski et al., 1989, 1990). Prestwich joined these efforts in the mid-1980s synthesizing valuable chemical probes and pheromone analogs (Prestwich et al., 1984; Vogt et al., 1985, 1988) and elucidating the pheromone binding properties of PBPs (Prestwich, 1985, 1987; Du et al., 1994; Du and Prestwich, 1995; Plettner et al., 2000; Lazar et al., 2002). Many more laboratories became interested in OBPs beginning in the late 1980s, greatly expanding knowledge of the range of species possessing these proteins and knowledge of the size and diversity of the OBP gene family in single species (reviews: Pelosi and Maida, 1995; Krieger and Breer, 1999; Vogt et al., 1999; Leal, 2003; NagnanLe Meillour and Jacquin-Joly, 2003; Plettner, 2003; Vogt, 2003). The transduction pathway was identified as G-protein/IP3-based during the early 1990s (Boekhoff et al., 1990a, 1990b, 1993; Breer et al., 1990; Riesgo-Esgovar et al., 1995; Krieger and Breer, 2003). ORs were finally identified from the sequenced genome database of Drosophila (Clyne et al., 1999; Gao and Chess, 1999; Vosshall et al., 1999). Gustatory receptors were subsequently identified in a similar manner (Clyne et al., 2000; Dunipace et al., 2001; Scott et al., 2001). Now, having recently crossed the technological watershed of genome sequencing, we can expect to see this information expand rapidly across the Insecta.

3.15.3. Transduction Events 3.15.3.1. Receptors

3.15.3.1.1. Among noninsect species, odor molecules are detected by G-protein-coupled receptors The view that odor molecules are detected by membrane-bound receptor proteins was still being debated as recently as the early 1980s, and references to insect sensilla being light guides for infrared radiation being emitted from oscillating odor molecules still appear occasionally. But one really must credit Lancet for his seminal efforts in the early to mid1980s, which firmly established the G-proteincoupled receptor (GPCR) model for odor detection. Working in vertebrates, Lancet and his group were the first to demonstrate odor-dependent increases in cAMP and the presence of G proteins in isolated olfactory neurons (Pace et al., 1985; Pace and

Lancet, 1986). Work that followed established the existence of both cAMP and IP3 based G-protein transduction pathways in vertebrate olfaction (Jones and Reed, 1989; Dhallan et al., 1990; Bakalyar and Reed, 1991), setting the stage for the identification of 7-transmembrane domain GPCR odor receptors in the rat (Buck and Axel, 1991). The following decade saw the identification of odor receptors in vertebrates from fish, amphibians, birds, and mammals (Freitag et al., 1998; Mombaerts, 1999). Vertebrate odor receptors comprise a large gene family, ranging from perhaps 100 members in fish to perhaps around 1300 in mammals (Zhang and Firestein, 2002; Young and Trask, 2002). These genes are quite conserved across the vertebrates; catfish receptors were initially identified using polymerase chain reaction (PCR) primers designed using rat OR sequences (Ngai et al., 1993), and a set of ‘‘universal’’ primers has been used to clone ORs from coelacanth to mammal (Freitag et al., 1998). Many details of this gene family, including its full size, are being revealed as full species genomes are made available. Interestingly, many OR genes have accumulated mutations rendering them nonfunctional pseudogenes, and the proportion of functional to nonfunctional genes varies considerably with the animal group and species (Frietag et al., 1998). For example, dolphins have no known functional OR genes (Frietag et al., 1998). Humans and mice have roughly similar number of OR genes, but in humans, nearly 70% are pseudogenes compared to perhaps 15% in mice (Young et al., 2002; Zhang and Firestein, 2002). The variation in functional members of this gene family can be considerable even among closely related species. Humans have as much as fourfold greater pseudogene content relative to other primates, a trend speculated to be related to differences in life history of the respective species (Rouquier et al., 2000; Gilad et al., 2003). Vertebrate pheromone receptors are also GPCRs. The vertebrate vomeronasal organ (VNO), known in reptiles and mammals, is a distinct olfactory epithelium whose neuronal axons project to a distinct olfactory bulb (Halpern, 1987; Halpern et al., 1998; Tirindelli et al., 1998; Dulac, 2000; Takami, 2002). Three classes of unique GPCR genes have been identified as VNO receptors in mice: V1R, 80 genes; V2R, 50–100 genes; V3R, 100–120 genes (Dulac and Axel, 1995; Herrada and Dulac, 1997; Matsunami and Buck, 1997; Ryba and Tirindelli, 1997; Pantages and Dulac, 2000; Martini et al., 2001; Zufall et al., 2002). The V2R class of VNO receptors has also been identified in fish (Cao et al., 1998; Naito et al., 1998), and a V2R from goldfish

Molecular Basis of Pheromone Detection in Insects

has been shown to interact with a variety of amino acids (Speca et al., 1999). The V2R receptors, at least, are sufficiently conserved across the vertebrates to have permitted fish V2R genes to be identified using PCR primers based on mammalian V2R sequences (Cao et al., 1998). An emerging view is that these VNO receptors may have originated as detectors of waterborne odor molecules, and have become increasingly specialized as pheromone receptors in higher vertebrates. Chemoreceptor proteins of the nematode Caenorhabditis elegans are also GPCRs (Troemel et al., 1995; Sengupta et al., 1996; Wes and Bargmann, 2001). Analysis of the fully sequenced C. elegans genome indicates the presence of several families of functional GPCR chemoreceptors totaling nearly 800 genes (Robertson, 2000), which is remarkable considering that C. elegans has only 16 pairs of chemosensory neurons (Bargmann and Mori, 1997). Many nonfunctional C. elegans chemoreceptor genes are present in the genome; one gene family (srh) includes 214 functional genes and 90 pseudogenes (Robertson, 2000). One major difference between the chemoreceptor proteins of vertebrate and C. elegans is that vertebrate OR genes are more or less expressed one allele per neuron (Chess et al., 1994) while C. elegans chemoreceptor genes may be expressed 40–50 per neuron (Robertson, 2000). Overall there seems to be remarkably little similarity between the chemoreceptor proteins of vertebrate and C. elegans, beyond the fact that they are GPCRs and they share sufficient structural features to suggest that both are 7-transmembrane domain proteins. Such extreme sequence divergence was discouraging to those hoping that common sequence domains between nematode and vertebrate OR genes would enable an expansion of these receptors to additional phyla, and suggests that rapid progress into other phyla may only be possible with the assistance of sequenced genomes. 3.15.3.1.2. Insect pheromones/odor molecules are detected by G-protein-coupled receptors The identification of GPCRs as odor receptors in vertebrates and nematodes (see Section 3.15.3.1.1), and the identification of a G-protein-coupled IP3 cascade in insect olfaction (see discussion below) set the stage for the identification of insect odor receptors in Drosophila. Efforts during the 1990s to identify ORs in a variety of insects using vertebrate or nematode OR sequences largely failed. One Drosophila gene that subsequently proved to be an OR was cloned by differential hybridization during the mid-1990s (DOR104/Or85e), but efforts to use this sequence to identify any additional genes were

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unsuccessful (Vosshall, 2003a). The problem was, and is, that it is difficult or impossible to recognize OR genes in an insect without the aid of the genome of that insect having been fully sequenced, obvious exceptions being close sibling species or recently duplicated gene pairs. Drosophila OR genes were identified from the partially sequenced genome, searching for any sequences that (1) were likely expressed, (2) encoded some degree of hydrophobicity, and (3) were otherwise unidentifiable. ORs emerged from the resulting pool, and were tentatively identified as such using histological methods (Clyne et al., 1999; Gao and Chess, 1999; Vosshall et al., 1999). Analyses of the more or less fully sequenced genome of Drosophila have identified at least 61 candidate OR genes (Dor) (Table 1) (Drosophila Odorant Receptor Nomenclature Committee, 2000; Hill et al., 2002; Vosshall, 2003a), and another 69 candidate GR genes (Dgr) a few of which are also likely involved in odor detection (Clyne et al., 2000; Dunipace et al., 2001; Scott et al., 2001; Hill et al., 2002); all appear to be GPCRs. The genome of A. gambiae contains at least 79 OR and 76 GR genes (Fox et al., 2001, 2002; Hill et al., 2002; Vosshall and Keller, 2003). Nine OR genes have been identified from a genome database of the moth Heliothis virescens, eight by comparison (BLAST) with the Dor sequences and one more from a low stringency cDNA library screen using the new H. virescens DNAs as probes; many more are presumably yet to be identified (Krieger et al., 2002). Sequencing efforts are currently being applied to the genomes of several insects of diverse orders, both closely and distantly related to those above, which will greatly expand our knowledge of the insect OR gene family, as well as other olfactory gene families. In adult Drosophila, olfactory neurons are restricted to the antennas (third segment) and maxillary palps. A total of about 1300 olfactory sensory neurons (OSNs) (1200 per antenna, 120 per palp) project axons to 43 glomeruli of the olfactory lobe (Venkatesh and Singh, 1984; Singh and Nayak, 1985; Stocker, 1994); all neurons expressing a given OR gene converge on one or in some cases a small subset of glomerulae (Gao et al., 2000; Vosshall et al., 2000; Bhalerao et al., 2003). Overall there is little difference between male and female antennas/ palps, though female antennas do have slightly more sensilla than males – 457 versus 419, primarily of the basiconic type (Shanbhag et al., 1999). Of the 61 Dor genes, 39 are expressed in antennal neurons and nine are expressed in palp neurons (Clyne et al., 1999; Gao and Chess, 1999; Vosshall et al.,

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Table 1 Drosophila olfactory receptor (OR) gene names and accession numbers Name a

Accession number

Or1a Or2a Or7a Or9a Or10a Or13a Or19a Or22ab Or22bb Or22c Or23a Or24a Or30a Or33ab Or33bb Or33cb Or35a Or42ab Or42bb Or43a Or43b Or45a Or45b Or46ab Or46bb Or47a Or47b Or49a Or49b Or56ab Or59ab Or59bb Or59cb Or63a Or65ab Or65bb Or65cb Or67a Or67b Or67c Or67d Or69ab Or69bb Or71a Or74a Or82a Or83ab Or83bb Or83c Or85a Or85bb Or85cb Or85d Or85e Or85f Or88a Or92a Or94ab Or94bb Or98a Or98b

CG17885 CG3206 CG10759 CG15302 CG17867 CG12697 CG18859 CG12193 CG4231 CG15377 CG9880 CG11767 CG13106 CG16960 CG16961 CG5006 CG17868 CG17250 CG12754 CG1854 CG17853 CG1978 CG12931 CG17849 CG17848 CG13225 CG13206 CG13158 CG17584 CG12501 CG9820 CG3569 CG17226 CG9969 NA NA NA CG12526 CG14176 CG14156 CG14157 NA CG17902 CG17871 CG13726 NA CG10612 CG10609 CG15581 CG7454 CG11735 CG17911 CG11742 CG9700 CG16755 CG14360 CG17916 CG17241 CG6679 CG5540 CG1867

1999, 2000); the remainder may be expressed at levels too low for the detection methods applied, or may express at specific developmental stages other than those examined, e.g., larval (Vosshall, 2003a), or may not be expressed at all. But clearly the majority of these OR genes are expressed and are apparently not pseudogenes. Each OSN is thought to express only one OR gene, though this is not fully confirmed; the one clear exception is Dor83b which actually appears to express in all olfactory neurons, coexpressing with other ORs (see below); on average, about 25 OSNs express any given OR gene (range 2–50 OSNs per OR gene) (Vosshall, 2003a). Dor genes encode proteins about 370–400 amino acids long; Dor83b is again an exception encoding a 486 amino acid protein. The presumptive 7-transmembrane domain of most Dor proteins contains a conserved motif Phe-Pro-XCys-Tyr-(X)20-Trp and the highest overall region of similarity among these proteins spans the 6- and 7-transmembrane domains (Vosshall, 2003a). The insect OR genes are highly divergent. In general, the Dor proteins show about 17–26% sequence identity, with a few more similar (40–60%) presumably due to recent gene duplications (Vosshall, 2003a). A comparison of the Anopheles and Drosophila OR genes indicates that the majority of these dipteran genes segregate by species, the only clear ortholog being Dor83b/AgamGPRor7 (Hill et al., 2002). OR genes of the moth H. virescens are similarly divergent, sharing very low sequence identity with the Dor proteins and only 7–16% identity with each other (Krieger et al., 2002). Analyses of all these proteins do suggest subgroupings based on sequence similarities (Hill et al., 2002; Krieger et al., 2002); whether these groupings reflect functional differences remains to be demonstrated. The Dor genes are distributed throughout the Drosophila genome. Most reside in isolated locations, though about one-third of the Dor genes reside in clusters of two or three, presumably deriving from related gene duplications (Table 1). One extreme example of such a duplication is the OR22a/22b pair. These two genes are separated by

a

Drosophila OR genes have been named after their map location. The Drosophila genome is artificially divided into 100 map regions: Chromosome 1 (C1), 1–20; C2, 21–60; C3, 61–99. Centromeres are located by map locations 40 and 80. b OR genes residing near enough to define a gene cluster originating from probable ‘‘recent’’ gene duplications. NA, not applicable. Modified from Vosshall, L.B., 2003a. Diversity and expression of odorant receptors in Drosophila. In: Blomquist, G.J., Vogt, R.G. (Eds.), Insect Pheromone Biochemistry and Molecular Biology. Academic Press, London, pp. 567–591.

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only about 650 bp, are 78% identical in their predicted amino acid sequences, and coexpress in the same OSNs (Dobritsa et al., 2003). The function of any of these genes as ORs is indirectly supported by their numbers, their divergent sequences, and their distributed expression among different OSNs (i.e., one OR per neuron). Dor43b has been immunologically localized to sensory dendrites (Elmore and Smith, 2001). Direct evidence that these genes encode ORs includes several electrophysiological studies. Ectopic expression of Dor43a in olfactory neurons using an OSN-specific driver gave increased responses in electroantennograms (EAGs) to several odor molecules including benzaldehyde, benzyl alcohol, cyclohexanol, and cyclohexanone (Sto¨ rtkuhl and Kettler, 2001). Coexpression of Dor43a and a Drosophila G protein in frog oocytes made these oocytes sensitive to these same odor molecules (but not other odor molecules tested, and sham-injected oocytes were nonresponsive), and provided support that the ORs were indeed G protein coupled (Wetzel et al., 2001). Dor22a, Dor22b, and Dor47a have also recently been characterized (Dobritsa et al., 2003). Dor22a and Dor22b are close gene neighbors (650 bp apart), share 78% amino acid sequence identity, and appear to coexpress in the same neuron in one class of basiconic sensilla (29 sensilla in female, 18 sensilla in male). Deletion/rescue electrophysiological studies suggest that Dor22a responds to ethyl butyrate, pentyl acetate, and ethyl acetate; however, no odor molecules (of 87 tested) were shown to stimulate Dor22b. This genetic system was shown to be useful for functionally characterizing other OR genes: expression of Dor47a using the Dor22a promoter and in a Dor22a/Dor22b null background demonstrated neuronal responsiveness to a set of odors novel to the Dor22a-expressing wild-type neuron. While these studies indicate that Dor22a and Dor22b coexpress, the only evidence presented that Dor22b might be functional is its identification in the sibling species D. simulans; observed conservation between the Dor22b orthologs in D. melanogaster and D. simulans was presumed possible only if selection was acting on the receptor genes, implying that the gene products are functionally active. Dor83b is an OR gene that stands apart from the rest: it is expressed in all OSNs, it encodes a protein that is structurally distinct from other Dor proteins, and it is the only OR gene readily identified in diverse species: the fruit fly D. melanogaster (Dor83b: Vosshall et al., 2000); the mosquito A. gambiae (AgamGPRor7: Hill et al., 2002), the

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moth H. virescens (HR2: Krieger et al., 2002), as well as the silk moths Bombyx mori and A. polyphemus, the honeybee Apis mellifera, the blowfly Calliphor erythrocephala, and the beetle Tenebrio molitor (Krieger et al., 2003). These 83b/GPRor7/ H2 proteins are 64–88% identical, differing from one another in a phylogenetically consistent manner (Krieger et al., 2003). As this gene expresses in all OSNs, always coexpressing with one of the other OR genes, it is presumably not functioning as the other OR proteins do in interacting with an odor ligand. Several functions for this gene have been speculated including that (1) it forms a heterodimer with the ligand-sensitive OR, (2) that it activates a second messenger pathway in a ligand independent manner, (3) that it helps in the trafficking of OR proteins to the correct site in the dendritic membranes (discussed by Krieger et al., 2003; Vosshall and Keller, 2003; Vosshall, 2003a). None of these functions has been verified as of early 2000s. Insect OR genes partially define the phenotypes of OSNs, but they apparently do not contribute to the determination of these phenotypes. Figure 1 summarizes general features of the brain neurocircuitry of the insect olfactory pathway and includes references to many elegant studies. Each glomerulus of the olfactory lobe receives axons belonging to OSNs of a single phenotype; all OSNs converging on an individual glomerulus are thought to express the same OR gene (Gao et al., 2000; Vosshall et al., 2000; Bhalerao et al., 2003). The positions of these glomerulae are more or less constant from animal to animal, and in some cases the preservation of these positions may have a functional significance in coding (Hansson et al., 1992, 2003; Carlsson and Hansson, 2003a, 2003b). Furthermore, a recent study has shown that the projection neurons emanating from these glomerulae terminate their axons in the lateral protocerebrum in a spatially restricted manner that to some degree maintains the spatial organization of the olfactory lobe (Wong et al., 2002). In vertebrates, OR genes have been shown to influence the projection of OSNs to specific glomerulae; neurons missexpressing the appropriate OR gene miss or fail altogether to target a specific glomerulus (Mombaerts et al., 1996; Wang et al., 1998; reviews: Korsching, 2002; Vosshall, 2003a, 2003b). The Drosophila ORs, however, apparently do not have this capability. OSNs expressing Dor22a/Dor22b target an identifiable glomerulus; these OSNs still target this glomerulus in a Dor22a/Dor22b deletion mutant whether or not an alternative OR gene is expressed in these neurons (Dobritsa et al., 2003). In insects, correct targeting of OSNs appears to function independent of the OR genes.

762 Molecular Basis of Pheromone Detection in Insects

3.15.3.1.3. Gustatory neurons detect chemostimulants using G-protein-coupled receptors Perhaps 69 GPCRs have been identified as candidate gustatory receptors in the Drosophila genome (Clyne et al., 2000; Dunipace et al., 2001; Scott et al., 2001; Hill et al., 2002); 76 were identified in the A. gambiae genome (Hill et al., 2002). The expression of a subset of the Drosophila GRs has been characterized; these are associated with a variety of tissues including the ventral pits, gut, mouth, dorsal, and terminal organs of larvae and labellum, cibarial organs, labral organ, legs, and wings of adults (Dunipace et al., 2001; Scott et al., 2001). One, Gr21D1, expresses in antennas, associating with a neuron that projects to a specific glomerulus in the olfactory lobe (Scott et al., 2001) However, all others examined which express in head tissues associate with neurons that project to the subesophageal ganglion (SEG) (Stocker, 1994). Like the antennal lobe, the SEG is organized into a discrete number of glomerulae, i.e., seven (Shanbhag and Singh, 1992). However, unlike the antennal system, GR expressing neurons arborize into multiple SEG glomerulae (Singh and Nayak, 1985; Singh, 1997; Dunipace et al., 2001; Scott et al., 2001). Only one GR gene has been functionally characterized; Gr5a encodes a trehalose receptor (Dahanukar et al., 2001; Ueno et al., 2001). Interestingly, in both studies Gr5a was chosen for study because it is tightly linked to another gene, tre (Ishimoto et al., 2000), which had previously been implicated as a trehalose receptor but has no sequence similarity to the putative Drosophila GR genes. Gr5a expresses in labellar sensilla. The GRs of Drosophila are highly divergent, ranging from 15% to 25% in sequence identity. Considering such divergence and the number of genes, these animals presumably ‘‘taste’’ a broad range of molecules that might further be quite different between larvae and adults. Insect taste is undoubtedly much more complex than may be implied by the understatement that taste discriminates the modalities of sweet, sour, salty, and bitter (Singh, 1997). As GRs are identified from a broad range of insects, they might be expected also to be highly divergent, driven so by the chemistry of host plants utilized by various insect groups. Whether any of these GRs are involved in pheromone detection remains to be determined. 3.15.3.2. Second Messenger Pathways

3.15.3.2.1. Pheromone/odor transduction is mediated by a G protein-based IP3 pathway The identification of GPCR odor receptors in both vertebrates and invertebrates was preceded by the identification of odor activated G protein/second messenger

cascades. Two G-protein-coupled transduction pathways have been identified in olfactory systems of vertebrates and arthropods: (1) G protein dependent activation of phospholipase C (PLC) with the subsequent production of IP3; or (2) G protein dependent activation of adenylate cyclase with the subsequent production of cAMP (Krieger and Breer, 2003). Both pathways exist in vertebrates, and in lobsters both pathways are thought to coexist in the same olfactory chemosensory neurons (Fadool and Ache, 1992; Michel and Ache, 1992; Boekhoff et al., 1994; Hatt and Ache, 1994). In insects, only the G-protein-coupled IP3 pathway illustrated in Figure 2c has been demonstrated for both pheromones and nonpheromonal odor molecules. When pheromone or other odor molecules were applied to homogenates of cockroach or moth antennas, a rapid rise in IP3 was observed (Boekhoff et al., 1990a, 1990b, 1993; Breer et al., 1990). This rise in IP3 was assumed to be mediated by a receptor activated G protein. The Ga subunits Go and Gq have been identified in or cloned from antennas of cockroach, fly, and moth (Breer et al., 1988; Boekhoff et al., 1990b; Raming et al., 1990; Talluri et al., 1995), and Gq-like proteins have been immunohistologically visualized in the dendritic membranes of pheromone sensory neurons in the silk moths B. mori and A. pernyi (Laue et al., 1997). To affect changes in IP3 levels, a receptor activated Ga subunit would activate the membrane-bound enzyme PLC, which in turn would cleave the membrane lipid phosphotidyl inositol 4,5-biphosphate (PIP2) to membrane-bound diacylglycerol (DAG) and cytosol soluble IP3 (see Figure 2c). A PLC Drosophila mutant, norpA, has been shown to be severely impaired in odor detection (Riesgo-Escovar et al., 1995). PLC has also been immunologically identified by Western blot in homogenates of olfactory dendrites of the moth A. polyphemus (Maida et al., 2000). The classic action of IP3 is to interact with Ca2þ channels, also referred to as IP3 receptors; such receptor/ channels have been immunohistologically visualized in the dendritic membranes of pheromone sensory neurons of B. mori and A. pernyi (Laue and Steinbrecht, 1997). In olfactory neurons of the locust Locusta migratoria, extracellular application of odors or intracellular application of IP3 generated an increase in the inward membrane conductance of cations, including Ca2þ (Wegener et al., 1997). In cultured adult olfactory neurons of the moth M. sexta, an IP3-dependent inward Ca2þ current was observed to precede general inward cation currents (Stengl, 1994). Thus, by 1999, all the components depicted in the Figure 2c scheme have been

Molecular Basis of Pheromone Detection in Insects

identified, with the exception of the odor receptors themselves. A limited number of studies have investigated the role of cAMP in insect olfactory transduction, but no increases in cAMP have been observed on stimulation with pheromone or other odor molecules in insects (Ziegelberger et al., 1990; Boekhoff et al., 1993). Curiously, cyclic nucleotide gated ion channels have been identified in insect antennas, and in fact is localized to olfactory receptor neurons (Baumann et al., 1994; Krieger et al., 1997). This may imply that a cAMP pathway does exist for certain odorants; cAMP olfactory pathways are known in lobsters (Michel and Ache, 1992; Hatt and Ache, 1994) and vertebrates (e.g., Jones and Reed, 1989; Bakalyar and Reed, 1990; Boekhoff and Breer, 1992; Rossler et al., 2000b). Alternatively, these cyclic nucleotide gated channels may function in other roles, perhaps as part of neuromodulatory pathways in the neuronal cell bodies, or in the maintenance of background neural activity and sensitivity (Krieger et al., 1997). The role of cGMP in pheromone transduction has also been demonstrated, but it appears to have a modulatory role in adaptation or desensitization rather than participating in primary transduction (see Section 3.15.3.2.2) (Ziegelberger et al., 1990; Redkozubov, 2000; Stengl et al., 2001; Dolzer et al., 2003). As of now, there is little evidence that cyclic nucleotides act in a primary capacity in the transduction of pheromone or other odor signals in insects. 3.15.3.2.2. Pheromone/odor transduction is modulated by G-protein based second messenger processes Second messenger processes have dynamic properties due to the number of divergent pathways they can initiate; few of these potential pathways have been investigated. These might be forward pathways, where signal molecules such as IP3 might interact with secondary targets, perhaps altering the future response characteristics of the neuron, or they might be backward pathways, downregulating the activity of processes just initiated (e.g., the receptor mediated G proteincoupled pathway). The physiological dynamics of pheromone adaptation has recently been investigated in the moth B. mori with respect to stimulus concentration and duration (Dolzer et al., 2003). Yet, few modulatory processes have been examined thoroughly in insect olfaction. Two loose ends depicted in Figure 2c are the bg subunits of the G-protein (Gbg) complex and DAG. ‘‘Gbg is now known to directly regulate as many different protein targets as the Ga subunit’’ (Clapham and

763

Neer, 1997). In vertebrates, Gbg has been shown to activate G-protein-coupled inwardly rectifying Kþ channels (GIRKs) (Salvador et al., 2003). Gbg has been shown to activate regulators of G-protein signaling (RGS) proteins which in turn activate GAP protein (GTPase activating protein) which regulates the GTPase activity in the Ga subunit (turning off Ga activity) (Witherow and Slepak, 2003). Gbg has been shown to regulate G-protein-coupled receptor kinases (GRKs) which phosphorylate and downregulate GPCRs (Inglese et al., 1992; Pitcher et al., 1998; Eichmann et al., 2003). Gbg has been shown to regulate the activities of certain PLC enzymes and the subsequent production of IP3 (Akgoz et al., 2002). In vertebrate chemoreception, Gbg has been shown to regulate PLC activity in mammalian vomeronasal neurons (Runnenburger et al., 2002) and in bitter taste neurons (Rossler et al., 2000a). Active Gbg signaling in insect olfaction has yet to be reported. Arrestins are proteins that bind to the phosphorylated state of a GPCR, and can mediate the uncoupling of GPCRs from G proteins (desensitization) (Inglese et al., 1993; Pippig et al., 1993; Freedman and Lefkowitz, 1996) as well as the internalization of GPCRs (with presumed subsequent dephosphorylation in microsomes with subsequent recycling of the receptor, i.e., resensitization) (Ferguson et al., 1996). In an IP3 mediated pathway, GPCR phosphorylation may be mediated by GIRKs, which may be mediated by Gbg (see above). Arrestins have been identified in insect antennas and shown to be critical in the olfactory transduction process in Drosophila and A. gambiae (Merrill et al., 2002). In this study, an arrestin was cloned from A. gambiae, and was shown to express in both olfactory and visual sensory neurons. Of three Drosophila arrestins, DmArr1 and DmArr2 were shown to express in both visual and olfactory tissue while DmKrz was nonvisual but also expressed in olfactory tissue. A double mutant (arr12; arr23) showed reduced EAG response to butanol and ethyl acetate, suggesting that arrestins play a key role in olfactory transduction in these animals (Merrill et al., 2002). DAG is another product of the IP3 pathway that has potential targets. DAG is classically considered as a regulator of protein kinase C (PKC), but may also interact with membrane lipases to induce the release of polyunsaturated fatty acids (e.g., arachidonic acid) which may activate certain ion channels/ membrane conductances. These ion channels may also be sensitive to levels of PIP2, regulated by PLC (Hardie, 2003; Krieger and Breer, 2003; Nilius et al., 2003). Recently, a membrane permeable analog of

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764 Molecular Basis of Pheromone Detection in Insects

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DAG, 1,2-dioctanol-sn-glycerol (DOG), was shown to elicit action potentials in pheromone sensitive neurons of the moth B. mori (Pophof, 2002; Pophof and van der Goes van Naters, 2002), suggesting an active role for DAG in insect olfactory transduction. Prolonged but only moderately elevated cGMP levels in response to applied sex pheromone have been measured in the antennas of the moths A. polyphemus and B. mori (Ziegelberger et al., 1990). Application of dibutyryl cGMP inhibited the response of B. mori pheromone sensitive neurons to pheromone (Redkozubov, 2000). These findings suggest a role for cGMP in modulating the response to sex pheromone. Pheromone-dependent elevation of cGMP levels was observed immunologically to occur in only a very small subpopulation of known pheromone sensitive sensilla (about 1% of sensilla trichoidea) of male antennas of the moth M. sexta (Stengl et al., 2001). In these same studies, the location of NO-sensitive soluble guanylate cyclase was determined to be entirely restricted to a population of sensilla chaetica but absent from sensilla trichoidea. Thus, while there appears to be a role for cGMP in pheromone transduction, the mechanism underlying that role or its value to pheromone perception remains unresolved. However, cGMP appears to regulate olfactory transduction in a subpopulation of rat olfactory neurons that have behavioral significance (review: Zufall and Munger, 2001), suggesting that perhaps the subpopulation of neurons in M. sexta which shows a pheromonedependent rise in cGMP may similarly serve a specialized behavioral function (Stengl et al., 2001).

3.15.4. Perireception Events Figure 2b summarizes a number of biochemical reactions that interact directly with pheromone and odor molecules in the extracellular space of the sensillum interior. Each sensillum contains at least three classes of protein with evolutionarily designed binding sites for these ligands: OBPs, ORs, and ODEs. Individual sensilla contain multiple members of each class, and the phenotypes of functionally distinct sensilla can be defined by the unique combinations of these gene products. These proteins comprise a biochemical network designed to process environmental signals. This network serves as a genetic interface between the animal and its environment, its components designed and organized by evolutionary selection; the single purpose of these gene products has allowed selection to proceed unconstrained from the need to balance requirements of other competing functions.

3.15.4.1. Odor Transport by Odorant Binding Proteins

Insect OBPs comprise a multigene family that includes a group of small hydrophilic proteins which bind and transport small hydrophobic ligands. Vertebrates also possess proteins that function as OBPs; however, vertebrate and insect OBPs are evolutionarily unrelated (Tegoni et al., 2000; Ramoni et al., 2001). The first insect OBP identified was the PBP of the silk moth A. polyphemus (Vogt and Riddiford, 1981). This 14 kDa protein appeared to be specific to the male antenna, was perhaps the most abundant soluble protein in the antenna, was located in the aqueous extracellular fluid that bathed the pheromone-sensitive neurons, and could bind sex pheromone. The concentration of ApolPBP within the sensillum fluid was estimated to be about 10 mM (Klein, 1987). Additional PBPs were subsequently identified by tissue specificity and N-terminal sequence from the moths Hyalophora cecropia, B. mori, Lymantria dispar, M. sexta, and Orgyia pseudosugata (Vogt, 1987; Vogt et al., 1989, 1991a); the first full length OBP sequences were obtained for M. sexta PBP1 (Gyo¨ rgyi et al., 1988) and A. polyphemus PBP1 (Raming et al., 1989). The identification of two PBPs in L. dispar was the first indication of multiple PBP genes within a single species (Vogt et al., 1989). The identification and cloning of the general odorant binding proteins GOBP1 and GOBP2 (Vogt and Lerner, 1989; Breer et al., 1990; Vogt et al., 1991a, 1991b) were the first indication that OBPs were members of a multigene family. These data on lepidopteran OBPs provided a basis for identifying the first OBPs in Drosophila (OS-E and OS-F: McKenna et al., 1994; PBPRP1-5: Pikielny et al., 1994; LUSH: Kim et al., 1998). Collectively, the OBPs described above might be viewed as the ‘‘gold standard’’ OBPs against which all others are compared. One hallmark feature of these proteins that is almost universally cited when presenting new OBP sequences is the positionally conserved presence of six cysteines (first noted by Breer et al., 1990). In more recent genomic analyses, the presence of these six cysteines plus a similar size and an acceptable result from BLAST analysis has been viewed as sufficient to consider these genes members of an OBP gene family (Galindo and Smith, 2001; Graham and Davies, 2002; Hekmat-Scafe et al., 2002). OBPs have now been identified in at least 41 species from orders throughout the Neopterous insects (Table 2; Figures 3 and 4), suggesting that they are common elements in olfactory sensilla of

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Molecular Basis of Pheromone Detection in Insects

Table 2 Odorant-binding protein (OBP) related proteins with published sequences

Species

Protein/gene name

Accession number a

Aseg-PBP Aper-PBP1 Aper-PBP2 Aper-PBP3 Aper-GOBP1 Aper-GOBP2 Aper-ABPX Apol-PBP1 Apol-PBP2 Apol-PBP3 Apol-GOBP2 Avel-PBP Bmor-PBP Bmor-GOBP1 Bmor-GOBP2 Bmor-ABPX Cfum-PBP

AH007940 X96773 X96860 AJ277265 Y10970 X96772 AJ002519 X17559 AJ277266 AJ277267 P34169 (peptide) AF177641 X94987 X94988 X94989 X94990 AF177643

Epos-PBP1 Epos-PBP2 Epos-GOBP2

AF416588 AF411459 AF411460 L41640

Lepidoptera Agrotis segetum Antheraea pernyi

Antheraea polyphemus

Argyrotaenia velutinana Bombyx mori

Choristoneura fumiferana Epiphyas postvittana

Galleria mellonella Heliothis virescens

Helicoverpa armigera

Helicoverpa zea Hyalophora cecropia Lymantria dispar

Mamestra brassicae

Manduca sexta

Orgyia pseudosugata Pectinophora gossypiella

Sericotropin (nonolfactory) Hvir-PBP Hvir-GOBP1 Hvir-GOBP2 Hvir-ABPX Harm-PBP Harm-GOBP1 Harm-GOBP2 Hzea-PBP Hcec-PBP Hcec-GOBP2 Ldis-PBP1 Ldis-PBP2 Ldis-GOBP2 Mbra-PBP1 Mbra-PBP2 Mbra-PBP4 Mbra-GOBP2 Msex-PBP1 Msex-PBP1 Msex-PBP2 Msex-PBP3 Msex-GOBP1 Msex-GOBP2 Msex-ABPX Msex-ABP1 Msex-ABP2 Msex-ABP3 Msex-ABP4 Msex-ABP5 Msex-ABP6 Msex-ABP7 Opse-PBP

X96861 X96862 X96863 AJ002518 AJ278992 AY049739 AJ278991 AF090191 P34175 (peptide) P34172 (peptide) AF007867 AF007868 P34173 (peptide) AF051143 AF051142 AF461143 AF051144 M21798 AF323972 AF117589 AF117581 M73797 AF323972 AF117577 AF117591 AF393491 AF393488 AF393490 AF393498 AF393499 AF393500 P34178 (peptide)

Pgos-PBP

AF177656

765

Table 2 Continued Accession number a

Species

Protein/gene name

Spodoptera exigua Synanthedon exitiosa Yponomeuta agnagellus

Spexi-GOBP2 Sexi-PBP Ycag-PBP

AJ294808 AF177660 AF177661

Acup-PBP1Acup-PBP2 Aoct-PBP1 Aoct-PBP2 Aosa-PBP1 Aosa-PBP2 Eori-PBP1 Eori-PBP2 Hpar-OBP1 Hpar-OBP2 Pdiv-OBP1 Pdiv-OBP2 Pjap-PRP1 Rpal-OBP2

AB040980 AB040141 AB040143 AB040981 AB040985 AF031492 AB040144 AB040986 AB026554 AB026556 AB026552 AB026553 AF031491 AF139912

Tmol-B1 Tmol-B2 Tmol-THP12

M97916 M97917 U24237

Coleoptera Anomala cuprea Anomala octiescostata Anomala osakana Exomala orientalis Holotrichia parallela Phyllopertha diversa Popillia japonica Rhynchophorus palmarum Tenebrio molitor

(nonolfactory) Diptera Aedes aegypti Anopheles gambiae

Aaeg-OBPRP Agam-OBP1 Agam-OBP2 Agam-OBP-1

AY062131 AF393487 AF393485 AF437884

(OS-F) Agam-OBP-2 Agam-OBP-3

AF437885 AF437886

(OS-E)

Ceratitis capitata

Culex quinquefasciatus

Agam-OBP-4 Agam-OBP-5 Agam-OBP-6 Agam-OBP-7 Agam-OBPRP-1 Agam-OBPRP-2 Agam-OBPRP-3 Agam-OBPRP-4 Agam-OBPRP-5 Agam-OBPRP-6 Agam-OBPRP-7 Agam-OBPRP-8 Agam-OBPRP-9 Agam-OBPRP-10 Agam-OBPRP-11 Agam-OBPRP-12 Agam-OBPRP-13 Agam-OBPRP-14 Agam-OBPRP-15 Agam-OBPRP-16 Ccap-OBPRP Ccap-OBPRP

serum protein serum protein Cqui-OBPRP

AF437887 AF437888 AF437889 AF437890 EAA00498 EAA00779 EAA00788 EAA00801 EAA01392 EAA01491 EAA03447 EAA03742 EAA03745 EAA06799 EAA06803 EAA07741 EAA07997 EAA09324 EAA12996 EAA14622 AJ252076 AJ252077 Y08954 Y19146 AF468212

Continued

766 Molecular Basis of Pheromone Detection in Insects

Table 2 Continued

Table 2 Continued

Species

Protein/gene name

Drosophila melanogaster

Dmel-99D Dmel-99C Dmel-99B Dmel-99A

Phormia regina

Dmel-93A Dmel-85A Dmel-84A Dmel-83F Dmel-83E Dmel-83D Dmel-83C Dmel-83B Dmel-83A Dmel-76A Dmel-69A Dmel-58A Dmel-58B Dmel-58C Dmel-58D Dmel-57E Dmel-57D Dmel-57C Dmel-57B Dmel-57A Dmel-56I Dmel-56H Dmel-56G Dmel-56F Dmel-56E Dmel-56D Dmel-56C Dmel-56B Dmel-56A Dmel-51A Dmel-50A Dmel-50B Dmel-50C Dmel-50D Dmel-50E Dmel-49A Dmel-47B Dmel-47A Dmel-46A Dmel-44A Dmel-28A Dmel-22A Dmel-19D Dmel-19C Dmel-19B Dmel-19A Dmel-18A Dmel-8A Preg-CSRBP (taste)

Accession number a

CG7584 CG12665 CG7592 CG18111 CG17284 CG11732 PBPRP4 CG15583 CG15583 CG15582 CG15582 OSE OSF LUSH PBPRP1 CG13517 CG13518 CG13524 CG13519 CG13429 AF457149 CG13421 AF457147 AF457148 AF457143 CG13874 CG13873 AF457146 CG8462 CG11218 CG15129a CG15129b CG11797 AF457145 ?? CG13940 ?? ?? CG13939 CG8769 CG13208 CG12944 CG12905 CG2297 PBPRP5 AF457144 PBPRP2 CG15457 CG1670 CG11748 CG15883 CG12665 S78710

Hemiptera Lygus lineolaris

Llin-LAP

AF091118

Lmad-PBP

AY116618

Dictyoptera Leucophaea maderae

(cockroach)

Accession number a

Species

Protein/gene name

Zootermopsis nevadensis (termite)

Znev-OBP1 Znev-OBP2 Znev-OBP3

AY135381 AY135382 AY135383

Amel-ASP1 Amel-ASP2 Amel-ASP4 Amel-ASP5 Amel-ASP6 S. amblychila S. aurea S. geminata S. globularia l. S. interrupta S. invicta (fireant)

AF166496 AF166497 AF393495 AF393497 AF393496 AF427889 AF427890 AF427905 AF427906 AF427891 AF459414

Hymenoptera Apis mellifera

(honeybee)

Solenopsis sp.

Gp-9 S. macdonaghi S. quinquecuspis S. richteri S. saevissima

AF427901 AF427902 AF427904 AF427892

Lmig OBP

AF542076

Orthoptera Locusta migratoria a

Accession numbers available as of August 2003.

at least 98% of all insect species (Vogt et al., 1999). OBPs have yet to be identified in preneopterous lineages (e.g., Odonata – dragonflies, Ephemeropotera – mayflies, Thysanura – silverfish, Archaeognatha – bristletails). Direct cloning and genome analyses have identified more than 50 OBP-related genes in Drosophila (McKenna et al., 1994; Pikielny et al., 1994; Kim et al., 1998; Galindo and Smith, 2001; Graham and Davies, 2002; Hekmat-Scafe et al., 2002; Vogt et al., 2002). A similar number have been identified in A. gambiae based on genome analyses (Robertson et al., 2001; Biessmann et al., 2002; Vogt, 2002; Xu et al., 2003). Most of these sequences are highly divergent, though some sequences are clearly the products of recent gene duplications based on sequence similarity, conserved exon structure, and clustered chromosomal location (e.g., Drosophila OS-E/OS-F: HekmatScafe et al., 2000, 2002; Vogt, 2002; Vogt et al., 2002). Like the Dors, the Drosophila OBPs are distributed throughout the genome (see Table 2, where DrosOBP genes are named for their map location) (Vogt, 2003). The majority of OBPs that have been sequenced in the Lepidoptera are PBPs and GOBPs which are distinct but share common sequence motifs; however, sequencing efforts have identified 13 OBP genes in M. sexta, including three PBPs, GOBP1, GOBP2, and another eight that are as highly divergent as the dipteran OBPs (Gyo¨ rgyi

Figure 3 A neighbor-joining tree of most OBPs listed in Table 2; bootstrap support is based on 1000 replicates and only branches with >50% support are shown. A single tree is shown, broken into thirds to fit the page. Insert (a) is a key to the taxa. Insert (b) is the uncollapsed tree (taxa not indicated) to illustrate full branch lengths (percent sequence differences). A size bar indicating 10% sequence difference is shown. Named branch clusters (e.g., ABPX) are referred to in the text.

768 Molecular Basis of Pheromone Detection in Insects

f0020

Figure 4 Overview of insect phylogeny (after Boudreaux, 1979; Hennig, 1981; Kristensen, 1991). In (a), the representation of a monophyletic orthopteroid group is considered debatable (Maddison and Maddison, 1998); this group was suggested by Boudreaux (1979) and Hennig (1981), but was not included by Kristensen (1991) due in part to uncertainties regarding the relationships of the Plecoptera. The monophyletic division of Endopterygota (holometabolous insects) (Kristensen, 1991; Whiting et al., 1997) is supported by most authors in part because of the unique development in this group; all members undergo a complete metamorphosis from the nonreproductive larval stage to the reproductive adult stage. A shared common ancestor for the Endopterygota and the hemipteroid lineages is suggested by morphological (Hennig, 1981; Kristensen, 1991; Whiting et al., 1997) and molecular data (Whiting et al., 1997). In (b), two supported endopterygote lineages are shown, one including the Coleoptera and a second including Hymenoptera, Diptera, and Lepidoptera. The fossil record suggests that the orthopteroids are more ancient than most endopterygote orders (late Carboniferous vs. early Permian); among the Endopterygota, the Coleoptera (Permian) are more ancient than the Diptera/Lepidoptera lineage, and the Diptera (Triassic) are more ancient than the Lepidoptera (Jurassic) (Kukalova´-Peck, 1991; Labandeira and Sepkoski, 1993). The greater part of molecular or biochemical based olfactory research has been done on a subset of these orders (noted by the insect symbols). Numbers of named species in each lineage are shown, taken from various entries in The Insects of Australia (Achterberg and CSIRO staff, 1991); these numbers are generally viewed as gross underestimates, but accurately reflect the relative success of the respective lineages.

et al., 1988; Vogt et al., 1991b, 2002; Robertson et al., 1999). Undoubtedly, full genome analysis of a lepidopteran species will identify many more. Not all OBPs are involved in odor detection. The genes listed in Table 2 include serum proteins from the medfly Ceratitis capitata (Y08954 and Y19146: Thymianou et al., 1998) and the beetle T. molitor (THP12: Rothemund et al., 1999; Graham et al., 2001); accessory gland proteins from T. molitor (B1 and B2: Paesen and Happ, 1995), and the Drosophila

protein PBPRP2 which is localized in nonchemosensory tissues or spaces (Park et al., 2000). Galindo and Smith (2001) examined the expression of 34 candidate Drosophila OBPs using a GAL4/UAS system, driving reporter gene expression using presumptive OBP promoters. Nine of these OBP-regulatory constructs expressed only in ‘‘chemosensory tissue’’ (sensilla of antennas and palps), four expressed only in ‘‘gustatory tissue’’ (sensilla of legs and wings), nine expressed in both tissues, five expressed

Molecular Basis of Pheromone Detection in Insects

in nonsensillar tissue, and seven did not express at all, perhaps due to inappropriate choice of regulatory region or the genes in question being pseudogenes (Galindo and Smith, 2001). Another recent study identified seven OBP-related genes that express in gustatory sensilla of the fleshfly Boettcherisca peregrina (Koganezawa and Shimada, 2002 (not included in Table 2). Clearly, though many OBP genes are assumed to express proteins which have an important role in odor detection, assigned membership in this gene family does not necessarily imply an olfactory or even chemosensory function; although, at the very least such membership should imply common ancestry (i.e., the genes all derived by gene duplications originating from one common ancestral gene).

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3.15.4.1.1. General properties of odorant binding proteins In general, OBPs are small, water-soluble, extracellular proteins around 14 kDa, and ranging between 120 and 150 amino acids long. OBPs presumably bind small ligands, and if involved in odor detection, those ligands are presumably odor molecules. OBPs are expressed with leader sequences which are removed during secretion. Most OBPs contain six cysteines in a certain asymmetric pattern, which has become a diagnostic feature of OBPs; these cysteines form disulfide bonds which stabilize the functionally relevant three-dimensional structure of the protein (Leal et al., 1999; Briand et al., 2001a, 2001b). OBPs involved in odor detection are expressed in sensilla support cells and secreted by these cells into the aqueous sensilla lumen (Vogt et al., 2002). The structure of OBPs has been characterized by X-ray crystallography and nuclear magnetic resonance (NMR) spectroscopy. X-ray analysis of the moth B. mori PBP1 (BmorPBP1) revealed six a-helix units stabilized by three disulfide bonds; a flask-shaped pheromone binding pocket is formed by four a-helices, and in an early study, another a-helical unit appeared to cover the narrow pocket opening (Sandler et al., 2000). Also, BmorPBP1 formed a dimeric structure in the crystal (also suggested by Leal, 2000). Similar structures have been determined for antennal OBPs from the honeybee and cockroach (Lartigue et al., 2003a, 2003b), as well as a hemolymph OBP homolog (THP12) of unknown function from the beetle T. molitor (Rothemund et al., 1999; Graham et al., 2001). Disulfide bridges join cysteines 1–3, 2–5, 4–6 in BmorPBP1 (Leal et al., 1999); the same disulfide bridge pattern has also been observed in the honeybee OBP ASP2 (Briand et al., 2001a, 2001b). Thus, even for OBP relatives with highly divergent

769

sequences, the overall structural features appear remarkably conserved. Additional structural information is presented below (see Section 3.15.4.1.3). 3.15.4.1.2. Differential expression of odorant binding proteins The unique expression of PBPs in antennas, their localization within olfactory sensilla, and their ability to bind sex pheromones was strong evidence of their role in pheromone detection (e.g., Vogt and Riddiford, 1981; Vogt et al., 1989). The identification of PBPs and GOBPs with their sexual dimorphic expression patterns suggested that different OBPs may associate with sensilla of distinct odor phenotypes (Vogt et al., 1991a). For example, in the moth M. sexta, PBP1 and GOBP2 express in different functional classes of olfactory sensilla (Vogt et al., 2002) (Figure 5). Initially, PBPs were thought to be male specific, but it is now clear that while PBPs associate with pheromone sensitive sensilla in males, PBPs also express in female antennas where their role is unknown. It is possible that PBPs expressing in females may function in the monitoring of pheromone release (Callahan et al., 2000; Vogt et al., 2002). Certain families of Lepidoptera, especially the noctuids, display an almost total sex indifference in antennal levels of PBP expression. The distribution of PBPs, GOBP1, GOBP2, and ABPX proteins have been characterized in several moth species using immunohistochemistry and electron microscopy; PBP proteins are present in the lumen of long trichoid sensilla, GOBP2 and ABPX proteins are present in the lumen of subsets of basiconic sensilla, and GOBP1 can be present in both trichoid and basiconic sensilla (Steinbrecht et al., 1995; Steinbrecht, 1996; Laue and Steinbrecht, 1997; Maida et al., 1999; Zhang et al., 2001). This broad expression of GOBP1 in both basiconic and trichoid sensilla was also observed by in situ hybridization in M. sexta (Vogt et al., 2002). Similar patterns have been described in Diptera. In Drosophila, OS-E, OS-F, and LUSH coexpress in trichoid sensilla, OS-E and OS-F but not LUSH coexpress in intermediate sensilla, and PBPRP5 expresses in a subset of basiconic sensilla (Hekmat-Scafe et al., 1997; Park et al., 2000; Shanbhag et al., 2001a). Galindo and Smith (2001) tested the expression patterns of 34 presumptive OBP promoters in Drosophila; nine drove expression in olfactory tissue alone (antenna and palps) while another nine drove expression in both olfactory and gustatory (wings and legs) tissues. These studies show that OBPs differentially express in sensilla that have diverse odor specificities, and therefore the OBPs contribute to the overall odor-specific phenotypes of the sensilla.

770 Molecular Basis of Pheromone Detection in Insects

Figure 5 In situ hybridizations (wholemount) of MsexPBP1, MsexGOBP2, and MsexGSTolf in adult male M. sexta antenna (a–e) and visualizations of sensory membrane proteins of Antheraea polyphemus, including ApolSNMP1 (f). Expression of M. sexta PBP1, GOBP2, and GST. (a) Shows the nonoverlapping distribution of pheromone sensitive long trichoid sensilla (i) vs. other olfactory sensilla (after Lee and Strausfeld, 1990). (b) and (c) show the nonoverlapping expression of MsexPBP1 and MsexGOBP2 (Vogt et al., 2002), and (d) shows the PBP-like expression of MsexGSTolf (Rogers et al., 1999). Arrows point to pheromone-sensitive sensilla. (e) shows a control in situ using an noninsect probe. Scale bar ¼ 100 mm. (Modified from Rogers, M.E., Jani, M.K., Vogt, R.G., 1999. An olfactory specific glutathione S-transferase in the sphinx moth Manduca sexta. J. Exp. Biol. 202, 1625–1637.) (f) Visualizations of A. polyphemus SNMP1. Three experiments characterizing proteins of the receptor membranes of olfactory neurons of male A. polyphemus. DZA: dendrite membranes were photolabeled with [3H]-DZA in the absence (P) and presence (+P) of excess nonradioactive pheromone; asterisk marks SHMP69 thought to be a candidate pheromone receptor (fluorogram, from Vogt et al., 1988). 35S-Met: dendrite membrane proteins metabolically labeled with injected 35S-Met, revealing a prominent band near 69 kDa (asterisk) as well as tubulin (T) and actin (a) (Vogt, unpublished fluorogram; for details of method see Vogt et al., 1989). Protein: membranes were isolated from olfactory sensilla isolated from 800 male antennas of animals attracted to female moths releasing sex pheromone. One-tenth of this preparation was analyzed revealing an abundant protein near 69 kDa (asterisk) visualized by Coomassie Blue staining; this band (ApolSNMP1) was electroblotted and sequenced (Rogers et al., 1997). Arrows identify the position of a 69 kDa marker in each gel.

p0225

p0230

3.15.4.1.3. Functional studies of odorant binding proteins The suggestion that OBPs function as odor transporters was first proposed (Vogt et al., 1985) as an alternative to a model that was prevalent at the time in which pore tubules served as the pheromone conduit from sensillum surface to OSN membrane (Ernst, 1969; Steinbrecht and Mu¨ ller, 1971; Kaissling, 1974, 1986; Kaissling and Thorson, 1980; Keil, 1982; Steinbrecht, 1997). Studies suggested that pore tubule to neuron contacts were quite rare (Keil, 1982), and their visualization highly dependent on tissue fixation in histological preparations (Steinbrecht and Mu¨ ller, 1971; Steinbrecht, 1980; Keil, 1982). In the likely absence of such a pore tubule delivery system, the presence of a soluble binding protein and a potent pheromone degrading enzyme suggested the model presented in Figure 2. In this new model, based on studies using the moth A. polyphemus, the hydrophobic pheromone molecules were solublized into the aqueous sensillum lumen by PBPs, transported by the PBPs to ORs in the OSN membranes, and subsequently degraded

by ODEs (Vogt et al., 1985). ODE kinetic properties were characterized, and the pheromone half-life within the sensillum was estimated at below 15 ms (Vogt et al., 1985). An even shorter half-life was later estimated for pheromone within sensilla of the moth M. sexta (Rybczynski et al., 1989). The ability of the A. polyphemus ODE to degrade pheromone was characterized at various pheromone concentrations and in the presence of various PBP concentrations (Vogt and Riddiford, 1986a). Results suggested that PBP did not protect pheromone from ODE through irreversible binding, but rather by mass action (i.e., PBP getting in the way of SE based on an estimated 1000-fold difference in concentrations for the two proteins). Results also suggested a pheromone-PBP KD in the 1–10 mM range (Vogt and Riddiford, 1986a). This function of PBPs as pheromone transporters in pheromone detection became generalized with the identification of PBPs from diverse species, and was extended to the detection of plant volatiles with the multispecies identification of GOBP1 and GOBP2 protein families (Vogt et al., 1991a). The

Molecular Basis of Pheromone Detection in Insects

identification of OBPs in Drosophila (McKenna et al., 1994; Pikielny et al., 1994) expanded these roles beyond the Lepidoptera; the identification of an OBP in the hemipteran Lygus lineolaris suggested that OBPs were present throughout the holometabolous and hemipteran lineages (Vogt et al., 1999), and the recent identification of OBPs in orthopteroid insects (Ishida et al., 2002a; Ban et al., 2003b; Rivie`re et al., 2003) argues that OBPs function throughout the neopteran lineages, and thus presumably the vast majority of eukaryotic organisms (Erwin, 1982; Novotny et al., 2002) (Figure 4). Finally, significant number of OBPs with divergent sequences have been identified in M. sexta, D. melanogaster, and A. gambiae (Table 2); such numbers are consistent with expectations for odor receptors, i.e., that there should be many odor receptors to deal with the number of odorous ligands an animal detects (Buck and Axel, 1991; Vosshall et al., 1999). The number and diversity of OBP genes (Table 2, Figure 3) coupled with their differential expression in olfactory sensilla during a time when odors are being detected (Figure 5) argues that they play a central role in odor processing. But what is that role? Pheromone binding has been demonstrated for PBPs from diverse species, including moths (Vogt and Riddiford, 1981; Vogt et al., 1989; Maı¨be`cheCoisne´ et al., 1997, 1998; Jacquin-Joly et al., 2000, 2001), honeybee (Danty et al., 1999), and beetle (Nikonov et al., 2002). Binding constants between pheromone and PBP have been estimated as low as 60 nM (Kaissling, 1986) but more typically in the range of 0.5–10 mM (Vogt and Riddiford, 1986a; Du and Prestwich 1995; Plettner et al., 2000; Kowcun et al., 2001; Campanacci et al., 2001a; Bette et al., 2002). Values of KD of 20–30 mM have also been determined for certain compounds and conditions (Kowcun et al., 2001; Bette et al., 2002); the KD for a PBP of Mamestra brassicae was estimated at 0.2 mM (Campanacci et al., 2001a). Two PBPs of the gypsy moth L. dispar were shown to differentially bind the two optical enantiomers of disparlure, the gypsy moth sex pheromone (Vogt et al., 1989; Plettner et al., 2000). Two PBPs of the silk moth A. pernyi were shown to differentially bind components of the sex pheromone of that species (Du et al., 1994; Du and Prestwich, 1995). Specific ligand binding has also been demonstrated for the GOBP2 protein of M. sexta (Feng and Prestwich, 1997) and a GOBP of the moth M. brassicae (Bohbot et al., 1998). A. polyphemus PBP1 (ApolPBP1) displayed different endogenous tryptophan based fluorescence in binding studies with three different pheromone components (Bette et al.,

771

2002). Differential specificities of the binding sites of PBPs of the moths M. brassicae and A. polyphemus have been demonstrated in competition studies examining the displacement of a fluorescent probe, 1-aminoanthracene (AMA) by pheromone components and nonpheromonal fatty acids (Campanacci et al., 2001a). One OBP from Drosophila, Lush, has been shown to be necessary for the appropriate behavioral response to alcohol; lush mutants do not display the normal wild-type avoidance of alcohol (Kim et al., 1998; Kim and Smith, 2001). Lush has also been shown to express in trichoid sensilla of the adult antenna, coexpressing with OS-E and OS-F (Shanbhag et al., 2001a). Nevertheless, for all these studies, the vast majority of OBPs remain uncharacterized, either biochemically or genetically. Pheromone binding may be regulated by pH dependent conformational changes in PBP. In BmorPBP1 the pheromone binding pocket is formed by four a-helices (Sandler et al., 2000). In crystals, a pheromone molecule interacts with diverse amino acids distributed throughout the protein but lining the binding pocket; both hydrophilic and electrostatic interactions are thus presumed to be critical for ligand binding specificity (Sandler et al., 2000; Klusa´k et al., 2003). It was initially suggested that an a-helix unit at the C-terminus covers the entry to the binding pocket and that this C-terminal a-helix must move to allow pheromone to enter or leave the binding pocket (Sandler et al., 2000). Solution NMR studies suggest that BmorPBP1 exists in two conformations which are pH dependent: a basic form (BmorPBPB) above pH 6.0 and an acidic form (BmorPBPA) below pH 4.9 (Damberger et al., 2000). More recent studies suggest that these conformational differences involve the C-terminus: at the higher pH range (i.e., physiological pH) the C-terminus is extended and lies on the surface of the protein (BmorPBPB) while at the lower pH range this same region forms a seventh a-helix which enters the binding pocket (BmorPBPA) and displaces a pheromone molecule bound within (Horst et al., 2001). A pH dependent pheromone binding was observed for the BmorPBP1; the pheromone–PBP complex was stable at pH 7.5 but significantly unstable at pH 5.5 (Wojtasek and Leal, 1999a). These studies have suggested that binding of pheromone at the pore tubule and release at the receptor may be driven by local pH differences within the sensillum, pH differences which alter the conformation of the PBP (i.e., from BmorPBPB to BmorPBPA) (Wojtasek and Leal, 1999a; Horst et al., 2001; Leal, 2003). The BmorPBPB conformation would favor binding of pheromone at the pore tubule and transporting it to the OR, while the

772 Molecular Basis of Pheromone Detection in Insects

BmorPBPA conformation would favor releasing pheromone to the OR. The proposed mechanism for such a pH gradient within a sensillum is the neuron membrane. The membrane phospholipids carry a negative charge (at the pH of interest) and are thus presumed to create a locally acidic boundary layer by attracting free hydrogen ions (actually, any cation); the conformation of the PBP–pheromone complex is thus changed from a bound state to a released state as it approaches the membrane (Wojtasek and Leal, 1999a). Other studies suggest alternative views regarding the mechanisms of odor–OBP binding. The crystal structure of an OBP from the cockroach Leucophaea maderae (LmadOBP) has recently been characterized in the presence of either the pheromone component 3-hydroxy-butan-2-one or the fluorescent molecule amino-naphthalen sulfonate (Lartigue et al., 2003b). This protein is missing the C-terminal region equivalent to the seventh a-helix in BmorPBP1, and therefore is missing the pH dependent opening/closing mechanism proposed for BmorPBP1 (see above). The binding pocket of the LmadOBP is also more hydrophilic than that of BmorPBP1. The authors suggest that these changes, especially the hydrophilic pocket, may be adaptive (Lartigue et al., 2003b), although LmadOBP is overall highly divergent from the other characterized OBPs. The lack of the C-terminal process in LmadOBP is especially interesting and brings to mind the presence/absence of the N-terminal inactivation structure of different voltage-gated ion channels. Clearly it will be helpful to evaluate and compare the structural properties of divergent OBPs from one species, as well as more OBPs from diverse species, in order to determine which features have functional significance and which merely reflect phylogenetic difference. Some studies also disagree with the pH-conformation hypothesis and to some degree with the crystallographic interpretations. The two PBPs of L. dispar show different KD for the two (þ and ) optical enantiomers of the pheromone (PBP1: KD(þ) 7 mM, KD() 2 mM; PBP2: KD(þ), 1.8 mM, KD(), 3 mM) (Plettner et al., 2000). However, this binding is not pH dependent over the ranges that would represent near-membrane conditions (pH 7.5 versus pH 5.5) (Kowcun et al., 2001). Modeling the membrane influence on the spatial distributions of Kþ and Hþ (i.e., pH) suggests that these influences only come into play at distances well under 2 nm from the cell membrane (Kowcun et al., 2001), which itself is only 8 nm thick. These studies suggest that pheromone offloading is not occurring by either pH or ionic strength gradients, but rather suggest that

the driving force to offload pheromone is either mass action kinetics (i.e., ORs have higher affinity for pheromone than PBPs) or direct interaction between the pheromone–PBP complex and OR (or perhaps some other membrane protein). These studies also observed an increased PBP binding capacity at high pheromone concentrations, and suggest that the PBP may form dimers dynamically under these conditions to trap pheromone at high concentrations to mitigate against stimulus saturation (Honson et al., 2003). The BmorPBP1 structural studies describe a deep binding pocket with a narrow opening covered by a C-terminal lid, and the involvement of a large numbered amino acids which interact with the majority of the pheromone molecule (Sandler et al., 2000; Klusa´ k et al., 2003). This seems a highly specific pocket and not terribly amenable to rapid release of pheromone; the conformational changes required would seem to require considerable energy, presumably provided by the proposed pH gradient, the presence of which has yet to be verified (it may be too shallow to have relevance, or it may be buffered by membrane proteins with high isoelectric points and thus be positively charged at pH 7.5). It was observed previously that PBP did not protect pheromone from degradation by a pheromone degrading esterase in vitro, though the PBP did maintain aqueous solubility of the pheromone, suggesting that a pheromone–PBP complex had formed (Vogt and Riddiford, 1986a). In contrast, the binding site described for BmorPBP1 would seem well designed for protection from enzymes. The binding site would also seem well designed for a very high degree of specificity, considering the number of interacting amino acid side chains (Klusa´ k et al., 2003). Yet studies examining the ability of various pheromone components and fatty acids to displace the fluorescent probe AMA suggest that the binding site may have relatively low specificity for its ligand(s) (Campanacci et al., 2001a). So, perhaps the binding site that is being described by crystallography is not the binding site used during rapid pheromone communication. And perhaps, though the PBPs may be conformationally sensitive to pH, these pH environments are not actually experienced in vivo. Clearly, one wants a system in which these hypotheses can be experimentally tested. Ideally one wants a genetic model, such as Drosophila, where OBPs can be modified or swapped and the result tested physiologically and/or behaviorally. 3.15.4.1.4. Evolutionary genomics of odorant binding proteins The OBP gene family was presumably derived through duplication of a gene with a func-

Molecular Basis of Pheromone Detection in Insects

tion that was likely not chemosensory, and the family has expanded through subsequent gene duplications. Alleles of these duplicated OBP genes accumulated, and evolutionary selection acted on these alleles leading to changes in the properties and functions of the predominant alleles in the species. Speciation events allowed allelic selection to proceed in manners supporting unique olfactory behaviors (life histories) of individual species. Depending on the function (nonchemosensory or chemosensory) of the founder gene(s), subsequent lineages diverged to either chemosensory or nonchemosensory functions. The genes currently listed as OBPs of D. melanogaster and A. gambiae include sequences with four to twelve cysteines (Hekmat-Scafe et al., 2002), differing significantly from the six cysteines of the gold standard OBPs, differences which assuredly influence the function of these proteins. Structural relationships between different OBPs can be explored to some extent using phylogenetic analysis tools (Figure 3). Such analyses compare amino acid sequences emphasizing similarities and differences. Amino acid sequences are aligned using programs such as ClustalX (Thompson et al., 1997), and trees are constructed using programs such as PAUP (Swofford, 2000) or MEGA (Kumar et al., 2001) and methods such as neighbor joining (Saitou and Nei, 1987) or maximum likelihood (Strimmer and von Haeseler, 1996; Nei and Kumar, 2000) available within these programs. The results of such analyses, when applied to proteins rather than nucleic acids, usually do not imply any evolutionary history, but rather suggest properties (characters) that certain proteins may share. Other criteria must be considered to infer evolutionary relatedness and history such as a demonstrated positive selection (e.g., significant nonsynonymous differences in codon nucleotides) or conservation of gene structure (e.g., exon–intron boundaries, relative positions in chromosomes, identification of neighboring genes: Vogt, 2002; Vogt et al., 2002). The sequence similarities of most OBPs listed in Table 2 are summarized in the neighbor joining tree illustrated in Figure 3. Overall, these OBPs are highly divergent, indicated by the relatively long lengths of most branches. Several branches include multiple taxa which define specific similarity groups; such groupings may suggest related functions. One such group is the Lepidoptera-specific PBP–GOBP gene family. Other similarity groups include a group of ant OBPs with a reputed role in governing the social behavior of Solenopsis spp. (fire ants and their relatives: Ross, 1997; Ross and Keller, 1998; Krieger and Ross, 2002), two groups that include the Drosophila OBPs LUSH and OS-E/OS-F, and a group

773

that includes the lepidopteran ABPX proteins. Both the LUSH and ABPX groups include genes belonging to multiple insect orders, a feature that may suggest that these proteins have a functional role common to or of central importance to diverse insects. However, most branches do not include OBPs from multiple orders, suggesting that selection acting on these genes is strongly biased by features unique to the orders, families, or genera represented. The PBPs and GOBPs of Lepidoptera represent a group that is not present in other insect orders, but these PBPs and GOBPs are clearly only a subgroup of lepidopteran OBPs. Their absence from Diptera, a sister lineage, and the relatively recent emergence of Lepidoptera implied by the fossil record, suggest that the PBP/GOBP genes may have arisen within Lepidoptera, or at least within the lepidopteran/ trichopteran lineage (Figure 4b). PBPs, GOBP1s, and GOBP2s comprise three distinct subgroups; amino acid sequence identities between PBPs and GOBPs are consistently between 25% and 30%, and amino acid sequence identities between GOBP1s and GOBP2 are consistently about 50%. GOBP1s and GOBP2s are highly conserved (around 90% identical within each group) while PBPs are highly divergent (20% to 95% identical). Also, multiple PBPs have been identified in individual species while only single genes are yet known for GOBP1s and GOBP2s, suggesting that gene duplications are occurring and being retained for PBPs but are either not occurring or not being retained for GOBPs. Clearly, selection is acting very differently on these subgroups, as might be expected for systems supporting reproduction (PBPs) versus general maintenance (i.e., feeding) (GOBPs). The genes encoding PBP1 and GOBP2 of M. sexta (MsexPBP1 and MsexGOBP2) are adjacent to one another, with coding regions separated by about 2000 bp, and both genes have identical exon–intron structures (Vogt et al., 2002). Proximity and conserved gene structure argue strongly that PBP1 and GOBP2 are derived from a gene duplication, yet the two proteins have very different sequences and temporal and spatial expression patterns. MsexPBP1 and MsexGOBP2 express in different types of sensilla (Figure 5a–c); MsexPBP1 expresses only in adult moths while MsexGOBP2 expresses in both larvae and adults, but the expression of both is regulated by ecdysteroid hormones (Vogt et al., 1993, 2002). If PBP1 and GOBP2 did derive from a gene duplication, then presumably GOBP1 derived from a subsequent duplication event involving GOBP2. The origins of multiple PBPs are less clear. Multiple PBPs may have been derived from

774 Molecular Basis of Pheromone Detection in Insects

subsequent duplications involving PBP1, or the PBP1/GOBP2 pair may have been derived from a duplication involving one of these other PBPs. PBP diversification may have been driven by reproductive selective pressures within species, genera, or families (Merrit et al., 1998); however, this does not appear to be the case with the GOBPs which are both highly conserved and broadly distributed among the lepidopteran families. An in-depth comparative study of the PBPs and GOBPs across the Lepidoptera might reveal much of the olfactory evolution of this insect order. OS-E and OS-F of Drosophila are also presumably derived from a gene duplication based on the neighboring relationship of their genes (separated by about 1000 bp), similar exon–intron structure, and similar sequence. However, unlike MsexPBP1/ MsexGOBP2 which differentially express, OS-E and OS-F coexpress in olfactory trichoid sensilla of the Drosophila antenna (Hekmat-Scafe et al., 1997); at least a subset of these sensilla contains neurons responsive to the pheromone cis-vaccenyl acetate (Clyne et al., 1997). Orthologs of OS-E and OS-F have been tentatively identified in A. gambiae, suggesting that the OS-E/OS-F gene duplication preceded the divergence of flies and mosquitoes within the Diptera (Vogt, 2002). A gene with significant sequence similarity to OS-E and OS-F has also been identified in the mosquito Culex quinquefasciatus (Ishida et al., 2002b); the sequence conservation of these presumed OS-E/OS-F orthologs among different groups of Diptera suggests their singular importance in the olfactory biology of these species. Unlike in Drosophila, the presumptive A. gambiae OS-E/OS-F genes are separated by about 30 million nucleotides suggesting that a chromosomal translocation event has occurred. It will be interesting to learn whether this separation has also resulted in a divergence in expression or function between these A. gambiae genes. The PBP/GOBP genes of Lepidoptera and OS-E/ OS-F genes of Diptera are just two stories of duplication and expansion of olfactory genes; the previously described pair of Drosophila OR genes, Dor22a and Dor22b, are another (Dobritsa et al., 2003) (see Section 3.15.3.1.2). A look at the distribution of OR, OBP, and SNMP homologs on the Drosophila chromosomes reveals a rich evolutionary history of gene duplication and translocation in the context of sensory biology and behavioral selection (Vogt, 2003). Increased access to genomes is permitting the identification of gene orthologs and the following of gene change across insect families and orders will allow us to hypothesize function in

one species based on the characterized function of a gene ortholog in another. 3.15.4.1.5. Allelic variation of pheromone binding proteins Several efforts have examined allelic variation of PBPs between distinct populations. Electrophoretic variants (possible multiple alleles) were described for both ApolPBP1 and ApolSE between populations of the moth A. polyphemus from New York (Long Island) and Wisconsin. However, this apparent allelic variation appeared high within, but similar between, each population and nothing definitive could be concluded regarding how this variation might affect behavior (Vogt, 1987; Vogt and Prestwich, 1988). LaForest and colleagues (1999) characterized PBP genes from individuals of two populations of the noctuid moth Agrotis segetum which use the same pheromone components but in different ratios. However, only neutral sequence variation was observed in the PBP sequences and again nothing definitive could be concluded regarding how such variation might affect pheromone perception and behavior. Similar results were obtained from studies of distinct populations of the European corn borer Ostrinia nubilalis (Willett and Harrison, 1999a, 1999b). Positive selection was observed for PBPs of species using different pheromone structures, but it was not clear that this selection resulted from the changes in pheromone components (Willett, 2000a, 2000b). A limitation of all three studies is that each characterized only one PBP; closer examination has demonstrated consistently that lepidopteran species have several PBPs (Table 2). It is likely that multiple PBPs derived from relatively recent gene duplications which occurred within specific lepidopteran lineages rather than at the base of the lepidopteran lineage (Merritt et al., 1998). If this is true, then it is also possible that, in certain species, PBP genes did not duplicate or that duplicated genes have become lost. But these possibilities are difficult to assess without a full genome sequence. If functionally relevant positive selection has occurred among the PBPs (which certainly seems likely), it may be discernable only by analyzing the larger repertoire of PBP genes within a species. 3.15.4.2. Odor Transport by OS-Ds, SAPs, or CSPs?

The proteins of another gene family, variously named OS-D, sensory appendage proteins (SAPs), or chemosensory proteins (CSPs), have been suggested frequently to have OBP-like properties in binding and transporting pheromones and other odors within sensilla (reviews: Picimbon, 2003;

Molecular Basis of Pheromone Detection in Insects

Nagnan-Le Meillour and Jacquin-Joly, 2003). These proteins have a wide tissue distribution throughout the body, share no sequence similarity to the OBPs, contain about 100–110 amino acid residues (compared with 140 for OBPs), four conserved cysteines (versus six in OBPs) and six a-helices (similar number but in a very different configuration than OBPs), and a hydrophilic channel that passes through the protein (versus a single-opening pocket in OBPs) (Campanacci et al., 2001b, 2001c, 2003; Briand et al., 2002; Lartigue et al., 2002). Table 3 lists members of this family reported to date. These proteins are remarkably conserved, showing consistently similar sequence identities independent of the phylogenetic distance of the organisms For example, CSPs share 50–60% identity between the two orthopteran species (Schistocerca gregaria and L. migratoria), 38–54% identity between the two lepidopteran groups CSPMbra A and CSPMbraB, and 37–50% identity between proteins of the Orthoptera versus Lepidoptera ( Jacquin-Joly et al., 2001). Such conservation is in stark contrast to the high degree of divergence observed for ORs and OBPs. Antennal CSPs were first identified through studies searching for antennal-specific genes. In their screen for antennal-specific proteins in Drosophila, Carlson’s group identified six cDNAs, naming them OS-A through OS-F (McKenna et al., 1994). This study yielded OS-E and OS-F, which were among the first OBPs identified outside of Lepidoptera (McKenna et al., 1994); one of the non-OBP proteins was named OS-D and was also described that year and named A10 by Pikielny in his characterization of Drosophila antennal proteins (Pikielny et al., 1994). Two years later, Pelosi’s group characterized antennal proteins of several species of phasmid (walking stick), identifying OS-D/A10 homologs in antenna and leg extracts of Eurycantha calcarata and Extatosoma tiaratum and antenna extracts of Carausius morosus (Mameli et al., 1996). Another 2 years later, Nagnan-Le Meillour’s group reported on the pheromone binding properties of antennal proteins in the moth M. brassicae (Bohbot et al., 1998), demonstrating nonspecific odor binding to a moth homolog of OS-D/A10 and noting its relationship to the phasmid proteins as well as proteins CLP1 from the labile palps of the moth Cactoblastis cactorum (Maleszka and Stange, 1997) and DA6b from the antenna of A. mellifera (Danty et al., 1998). In 1999, Robertson’s group reported their characterization of hundreds of antennal cDNAs of the moth M. sexta, including

775

Table 3 Chemosensory protein (CSP) related proteins with published sequences

Species

Protein/ gene name

Accession number a

BmorCSP1 BmorCSP2 Ccac CLP1 HarmCSP HvirCSP1 HvirCSP2 HvirCSP3 CSPMbraA1 CSPMbraA2 CSPMbraA4 CSPMbraA5 CSPMbraA6/A3 CSPMbraB1 CSPMbraB2 CSPMbraB3 CSPMbraB4 MsecSAP1 MsecSAP2 MsecSAP3 MsecSAP4 MsecSAP5

AAM34276 AAM34275 AAC47827 AAK53762 AAM77041 AAM77040 AAM77042 AAF19647 AAF19648 AAF19650 AAF19651 AAF71289(A6) AAF19652 AAF19653 AAF71290 AAF71291 AAF16696 AAF16714 AAF16707 AAF16721 AAF16716

LhumCSP

AAN01363

agCG50175 agCG50200 agCG50208 agCG50210 agCG50220 agCP11484 AgSAP1 DmelOS-D/A10 CG30172 CG9358 PEBmeIII RH74005/ CG11390

EAA12703 EAA12591 EAA12601 EAA12338 EAA12353 EAA12322 AAL84186 AAA21358 (OS-D) AAM68292 AAF47307 AAA87058 AAM29645 (RH)

LmigOS-D1 LmigOS-D2 LmigOS-D3 LmigOS-D4 LmigOS-D5 p10

CAB65177 CAB65178 CAB65179 CAB65180 CAB65181 AAB84283

CSPsg1 CSPsg2 CSPsg3 CSPsg4 CSPsg5

AAC25399 AAC25400 AAC25401 AAC25402 AAC25403

CSPec1 CSPec2 CSPec3

AAD30550 AAD30551 AAD30552

Lepidoptera Bombyx mori Cactoblastis cactorum Helicoverpa armigera Heliothis virescens

Mamestra brassicae

Manduca sexta

Hymenoptera Linepithema humile (ant)

Diptera Anopheles gambiae

Drosophila melanogaster

Dictyoptera Locusta migratoria

Periplaneta americana Schistocerca gregaria

Phasmatodea Eurycantha calcarata

a

Accession numbers available as of August 2003.

776 Molecular Basis of Pheromone Detection in Insects

five OS-D/A10 homologs which they named as sensory appendage proteins (SAPs) (Robertson et al., 1999). In 2000, Pelosi’s group characterized five OS-D/A10 homologs from the locust S. gregaria, referring to them as chemosensory proteins (CSPs) (Marchese et al., 2000). Table 3 lists the members of this gene family reported to date. CSPs associate with a broad range of body tissues, including legs, heads, thorax, antennas, proboscis, subcuticular epithelium, pheromone glands, and ejaculatory duct (Nomura et al., 1992; Dyanov and Dzitoeva, 1995; Nagnan-Le Meillour et al., 2000; Picimbon et al., 2000a, 2001; Jacquin-Joly et al., 2001; Picone et al., 2001; Ban et al., 2003a). CSPs may be absent from the central nervous system (Picimbon et al., 2000b). The name CSP (or SAP) is optimistically biased towards the chemosensory functions of these tissues, though clearly each tissue performs additional functions. Picimbon et al. (2000a, 2001) found that CSPs of the moths B. mori and H. virescens express in antennas, proboscis, legs, thorax, and head. The earliest identified member of the family, P10 of the cockroach Periplaneta americana, also expresses in legs, antennas, and heads (Nomura et al., 1992; Kitabayashi et al., 1998). In H. virescens, CSP expression in legs is initiated at least 5 days before adult eclosion, during a period when development is still ongoing, and expression persists into adult life; however, this time course does not match that of proteins where specific olfactory association is well established, such as OBPs and SNMP which tend to initiate expression only shortly before adult eclosion (Vogt et al., 1989, 1993; Rogers et al., 1997, 2001b; Picimbon et al., 2001). In the cockroach, P10 expression was primarily associated with leg regeneration (Nomura et al., 1992). While it may be true that most external insect tissues contain chemosensory organs, the mere association of CSPs with these tissues does not necessarily correlate with chemosensory function. Several CSPs have been shown to associate with chemosensory sensilla, though not in a consistent pattern. In Drosophila, OS-D/A10 expression (in situ hybridization) was associated with the sacculus, an antennal structure rich in coeloconic sensilla, although actual association with these sensilla was not shown (Pikielny et al., 1994). In the locust S. gregaria, antisera to CSPs purified from tarsi reacted with antigen in the lumen of gustatory sensilla of legs, palps, and antennas (Angeli et al., 1999). In the moth M. brassicae, CSPMbraA6 expression (in situ hybridization) was associated with hair-like olfactory sensilla (trichogen or

basiconic) but not with coeloconic sensilla (Jacquin-Joly et al., 2001). Several ligand binding studies have been reported for these proteins. CSPs from M. brassicae have been shown to interact with a wide range of hydrocarbons ranging in length from C9 to C18 and containing a variety of functional groups; KD values for all of these molecules are close to 1 mM (Bohbot et al., 1998; Jacquin-Joly et al., 2001; Lartigue et al., 2002; Campanacci et al., 2003; Mosbah et al., 2003). CSPs may be capable of receiving multiple ligands within their binding pocket; CSPMbraA6 was shown to bind ligands with a stoichiometry of 3 : 1 (Campanacci et al., 2003). A CSP from the honeybee was shown to interact with C14–C18 compounds, including several compounds reported to be brood pheromones, with KD values in the 1 mM range (Briand et al., 2002). A broad range of ligands was observed to interact with a CSP from the locust S. gregaria in competition studies with the fluorochrome N-phenyl-1-naphthylamine; binding of the fluorochrome was heat stable (100  C, 20 min) and had a KD of 4 mM, and KD values of competitors ranged from 10 to 150 mM (Ban et al., 2002, 2003b) (such heat stability in a soluble, globular protein with a-helices and disulfide bonds seems somewhat surprising). CSPs from the wing of the locust L. migratoria were purified with the oleoamide bound to the protein, a compound interpreted as an endogenous ligand for this CSP (Ban et al., 2003a); oleoamide is a C18 compound used as an industrial lubricant (recombinant PBPs have been isolated bound with spurious fatty acids) (Oldham et al., 2001). Presumably addressing the remarkable lack of discrimination of these proteins, Lartigue et al. (2002) wrote ‘‘We hypothesize therefore that CSPMbraA6 should interact with membranes to extract pheromones or other lipidic compounds dispersed in the hydrophobic membrane matrix and bring them to their specific target.’’ It does appear that at least some CSPs associate with chemosensory sensilla, and that they are capable of binding odor-like molecules, if with relatively low selectivity. And their presence in a broad range of insects and tissues together with their small ligand carrying capacity marks them proteins of potentially significant physiological interest. Nevertheless, the importance of these proteins in chemodetection remains remarkably unclear. 3.15.4.3. Pheromone and Odor Degradation by Odor Degrading Enzymes

Signal termination plays a critical role in all chemically mediated biological processes, and this is no

Molecular Basis of Pheromone Detection in Insects

less so in odor detection. The process of pheromone degradation has been studied for some years, at least as far back as Kasang (1971) in B. mori and Ferkovich et al. (1973a, 1973b) in the cabbage looper moth, Trichoplusia ni. Since then, a few pheromone degrading enzymes have been identified and characterized in detail and the general principal of pheromone degradation has become well established. Yet surprisingly little work is being done to investigate odor degrading enzymes (ODEs), in contrast with current efforts on ORs and OBPs. One reason to expand efforts on ODEs is their potential in insect control. If it is true that pheromones are perceived as precise mixtures, then the targeted inhibition of the ODE for a specific component should alter the blend ratio within a sensillum resulting in misperception of the pheromone. Of the three protein classes (ORs, OBPs, and ODEs), ODEs may be the least specific and thus the more generally targetable protein for behavioral inhibition. ODEs are enzymes selectively evolved to degrade odor molecules (Prestwich, 1987). However, to call an enzyme an ODE it is not enough to merely demonstrate the ability of an enzyme to degrade an odor; that enzyme has to be shown to reside in a space that is relevant to odor detection. This principle should be applied to OBPs and ORs as well. In general, those ODEs studied have been ones that attack specific functional groups, such as acetate esters, aldehydes, alcohols, ketones, and epoxides. ODEs attacking such different functional groups presumably belong to different gene families. The 55 kDa antennal-specific esterase (ApolSE) (see Section 3.15.4.3.1.1), that degrades the acetate-ester pheromone component of the silk moth A. polyphemus (Vogt and Riddiford, 1981; Vogt et al., 1985), is a member of a gene family that encodes insect esterases, an assumption supported by the recent cloning of a candidate ApolSE using PCR primers based on conserved regions of insect esterase genes (Ishida and Leal, 2002a). Similarly, the 150 kDa antennal-specific AOXs that degrades the aldehyde components of the A. polyphemus and M. sexta pheromones (Rybczynski et al., 1989, 1990) must belong to a distinct gene family of insect AOXs. So far, only the antennal-specific ApolSE and glutathione-S-transferase (MsexGSTolf) have been cloned (Rogers et al., 1999; Ishida and Leal, 2002). The following section organizes ODEs somewhat artificially into four categories, based on their location in the animal. To some degree this has functional significance, distinguishing between ODEs of the sensillum lumen (soluble or membrane bound), support cell cytosol and body surface.

777

3.15.4.3.1. Category 1: soluble extracellular ODEs of the sensillum lumen The early studies of the fate of pheromone within the antenna were among the first biochemical studies of insect odor detection. At the time, much seemed to be known about pheromone detection in moths: the pheromone of B. mori had been identified and radiolabeled (Butenandt et al., 1959; Kasang, 1968); pheromone detection had been characterized by electrophysiology in B. mori (Schneider, 1969); the ultrastructure of pheromone sensitive sensilla had been described for B. mori (Steinbrecht and Mu¨ ller, 1971). Adam and Delbru¨ ck (1968) had used the B. mori pheromone system to justify their ‘‘reduction in dimension’’ hypothesis for ligand-receptor targeting. Steinbrecht and Mu¨ ller (1971) had suggested that pore tubules might serve as conductive structures for the transport of pheromone molecules from the sensillum surface to receptors of the olfactory neurons (this view was later challenged, by Vogt et al. (1985), with the suggestion that OBPs performed odor transport). With this background, a series of studies were performed where radiolabled bombykol was applied to intact male B. mori antennas and the products analyzed following solvent extraction. These studies observed the general degradation of the alcohol bombykol to its acid metabolite, but identified no specific enzymes or enzyme activities (Kasang, 1971, 1973; Kasang and Kaissling, 1972; Kasang and Weiss, 1974; Kasang et al., 1989a, 1989b). About the same time an independent series of studies examined degradation of the T. ni pheromone from the active acetate ester to the inactive alcohol metabolite; these studies demonstrated esterase activities in the antenna but did not identify any antennal-specific enzymes (Ferkovich et al., 1973a, 1973b, 1980, 1982; Mayer, 1975; Taylor et al., 1981). Although no specific enzymes were identified, these efforts drew attention to the issues of pheromone degradation as an important component of the pheromone detection process. 3.15.4.3.1.1. Antennal-specific esterase The antennal-specific sensilla esterase (SE) of the moth A. polyphemus (ApolSE) was the first identified pheromone degrading enzyme (Vogt and Riddiford, 1981). The activity of ApolSE was visualized on nondenaturing polyacrylamide gel electrophoresis (PAGE), and was shown to be specific to the male antennas and to be located in the sensillar fluid (Vogt and Riddiford, 1981). ApolSE molecular mass was estimated at about 55 kDa (Klein, 1987). Antheraea polyphyemus was reported to have a two-component pheromone consisting of 9 : 1 ratio of acetate : aldehyde (Kochansky et al., 1975).

778 Molecular Basis of Pheromone Detection in Insects

ApolSE was shown to degrade the acetate component and was subsequently purified and subjected to kinetic analysis; a spectrophotometric assay was established using a- and b-naphthyl acetate as substrates, and a radioactive thin-layer chromatography (TLC) assay was established using tritiated pheromone (Vogt et al., 1985). ApolSE strongly preferred b- over a-naphthyl acetate, was inhibited by the volatile trifluoroketone 1,1,1-trifluoro-2-tetradecanone (IC50 ¼ 5 nM), and, most importantly, degraded the sex pheromone with unexpected aggression. By making adjustments for the concentrations and volumes within a sensillum lumen, the in vivo half-life of pheromone was conservatively estimated to be 15 ms in the presence of this enzyme. The observation of rapid pheromone degradation prompted the proposal of a new model for pheromone reception, one in which OBPs served as pheromone transporters (replacing pore tubules in that role), and enzymes degraded pheromone molecules to rapidly terminate these odor signals within the sensillum (see Figures 2 and 8). ApolSE was used to assess interactions between pheromone and PBP (Vogt and Riddiford, 1986a). ApolSE and ApolPBP1 were purified from antennas, and the ability of a fixed concentration of ApolSE to degrade pheromone was examined in various concentrations of PBP1 and pheromone. ApolSE activity was unaffected under conditions where PBP1 concentrations exceeded pheromone concentrations by 1000 times, suggesting that the PBP did not provide specific protection to pheromone from ApolSE, and did not bind pheromone irreversibly. Reduced activity was observed at all pheromone concentrations when PBP1 exceeded 1 mM. It was suggested that this transition was a consequence of protection during binding site-occupancy, but that occupancy must be transient, and that the KD of the pheromone–PBP interaction must be in the range of 1–10 mM. The activity of ApolSE below 1 mM PBP further suggested that pheromone–PBP binding was either very slow or very ephemeral (binding and release occurred rapidly). These conclusions are currently being challenged by alternative views that the pheromone–PBP complex may be quite stable, requiring some secondary interaction to drive dissociation (see earlier discussions regarding OBP function). An antennal-specific esterase was recently cloned from A. polyphemus and tentatively identified as ApolSE (accession no. AY091503) (Ishida and Leal, 2002). PCR primers were designed to conserved regions of other known insect esterases, and successfully used to amplify this cDNA. The cDNA encodes a protein with a predicted mass of

59 994 Da (553 amino acids, pI of 6.63) and three potential N-glycosylation sites. PCR studies confirmed that mRNA expression is male antenna specific. Kinetic analysis of expressed enzyme and histological localization should confirm the identity of this enzyme, but have not yet been reported. The apparent verification that pheromone degrading esterases are so readily identifiable, and the large number of species utilizing acetate ester pheromones suggest that a door has recently been opened wide for the future characterization of pheromone degrading esterases. 3.15.4.3.1.2. Antennal-specific aldehyde oxidase An antennal-specific aldehyde oxidase (AOX) of the moth M. sexta (MsexAOX) was the next identified pheromone degrading enzyme (Rybczynski et al., 1989). The activity of MsexAOX was visualized on nondenaturing PAGE, and was shown to be antennal specific but present in sensilla of both male and female antennas. MsexAOX was observed as a dimer with a combined estimated molecular mass of 295 kDa. Manduca sexta uses a multicomponent pheromone consisting exclusively of aldehydes including bombykal (Starratt et al., 1979; Tumlinson et al., 1989, 1994); MsexAOX was shown to degrade bombykal to its carboxylic acid, in the absence of any additional cofactors. Both TLC and spectrophotometric assays were established and a variety of substrates and inhibitors were characterized. Making adjustments for the concentrations and volumes within a sensillum lumen, the in vivo half-life of pheromone was estimated at 0.6 ms in the presence of this enzyme (Rybczynski et al., 1989). Many species use pheromone components with chemically diverse functional groups, and thus presumably require multiple ODEs. The pheromone of A. polyphemus is a blend of acetate and aldehyde components (Kochansky et al., 1975), and that of B. mori is a blend of alcohol and aldehyde (Kasang et al., 1978). Both A. polyphemus and B. mori have antennal-specific AOXs; both proteins were identified in antennal extracts, and ApolAOX was identified in extracts of isolated pheromone sensilla of male antennas (Rybczynski et al., 1990). ApolAOX and BmorAOX are both high molecular weight proteins, similar to MsexAOX, and are present in male and female antennas. Both ApolAOX and BmorAOX can degrade bombykal in the absence of any additional cofactors (Rybczynski et al., 1990). Bombyx mori antennas thus contain both alcohol oxidase or dehydrogenase activity (Kasang et al., 1989b) and antennal-specific AOX activities, and

Molecular Basis of Pheromone Detection in Insects

A. polyphemus sensilla contain both antennalspecific SE and AOX activities (Rybczynski et al., 1990). ApolSE is only found in male antennas, while ApolAOX is abundant in both male and female antennas, suggesting that ApolSE was selected to function specifically in pheromone degradation while ApolAOX (and presumably MsexAOX and BmorAOX) has the multiple functions of inactivating both pheromone and plant volatile odorants (Rybczynski et al., 1990). Tasayco and Prestwich (1990a, 1990b, 1990c) studied AOX and aldehyde dehydrogenase (ALDH) activities in the antennas of noctuid moths Heliothis virescens, H. subflexa, and Helicoverpa zea. Antennal-specific AOXs were observed in both males and females; molecular weights were similar to those of M. sexta, A. polyphemus, and B. mori. The noctuid enzymes also converted aldehyde pheromone components to their corresponding carboxylic acid without the aid of additional cofactors. Antennnal-specific ALDH enzymes were detected through radiolabeling with a substrate analog, but the location of the enzymes within the antenna was not determined. These ALDH enzymes required the cofactor NAD(Pþ). ApolSE and ApolAOX, as well as the other AOXs described above, function without the assistance of any cofactors, enabling them to act autonomously in the extracellular environment of the sensillum lumen where access to cellular metabolites might be limited. The ALDH requirement for cofactors may suggest that these enzymes are located in the cytoplasm of support cells where cofactors would be available. Antennal ALDH may serve as a biotransformation enzyme for pheromones and possibly xenobiotics in a manner similar to the antennal-specific GST of M. sexta (Rogers et al., 1999) (see Section 3.15.4.3.3). Several studies have reported slow rates of pheromone degradation on antennas (Kasang, 1971, 1974; Kasang and Kaissling, 1972; Kanaujia and Kaissling, 1985; Klun et al., 1991, 1996, 1998). These studies typically applied high concentrations of pheromone to antennas and subsequently extracted metabolites by incubating treated intact antennas in solvent, and analyzed the resulting solvent extract. It is difficult to interpret many of these results as no efforts were made to identify the tissue location of observed enzymatic activity. Enzymes capable of degrading pheromone are present in the cuticle, the cytoplasm of cells, and the hemolymph, in addition to the lumen of sensilla; observed activities may have been from any of these tissue spaces. It was suggested previously that observed slow degradation rates may be due to significant portions of pheromone migrating into the cuticle matrix rather than directly

779

entering sensilla, and that pheromone thus entering the cuticle matrix might exit the cuticle and encounter enzymes at slow rates and thus be degraded at slow rates (Vogt et al., 1985; Vogt, 1987). 3.15.4.3.2. Category 2: membrane-bound ODEs While it might seem most efficient for ODEs to be soluble and evenly distributed throughout the aqueous sensillum lumen, certain classes of enzymes may only be available for selection in membrane-bound forms. Such may be the case with pheromone degrading epoxide hydrolases (EH). The epoxide pheromone disparlure can be degraded by EH activity that is antennal specific and associated with isolated sensilla; this activity is not water soluble, suggesting that it might instead be associated with neuronal or support cell membranes (Prestwich, 1987; Vogt, 1987; Graham and Prestwich, 1992, 1994). Similarly, but in a noninsect system, a membrane-bound ODE that degrades purinergic odor molecules has been characterized in the spiny lobster, Panulirus argus (Trapido-Rosenthal et al., 1987, 1990; Gru¨ nert and Ache, 1988; Carr et al., 1990; Gleeson et al., 1992). Lobsters, arthropods by all rights, smell many things, including the purines ATP and ADP; the ratio of these molecules is thought to inform the lobster of how recently its food was alive (Zimmer-Faust et al., 1988). Certain olfactory neurons are stimulated by ATP and ADP, and the membranes of these neurons contain enzymes which degrade these purinergic signals (Trapido-Rosenthal et al., 1987, 1990; Gleeson et al., 1992). These enzymes are localized to membrane regions at the base of the sensillum, with their activities oriented towards the lymph cavity. 3.15.4.3.3. Category 3: Cytosolic ODEs – multifunctional ODEs inactivating odors and xenobiotics Olfactory and respiratory tissues of animals are constantly under attack by toxic chemicals from the environment. These xenobiotic compounds have presumably driven the enrichment in such tissues of biotransformation enzymes such as glutathionine-Stransferases (GSTs), cytochrome P-450 oxygenases, and UDP-glucuronosyltransferases, and various dehydrogenases and oxidases (reviews: Dahl, 1988; Mannervik and Danielson, 1988; Clark, 1990; Feyereisen, 1999). Many of these enzymes can also attack odor molecules, and this has led to suggestions for mammals that biotransformation molecules may function in odor signal termination (Nef et al., 1989; Lazard et al., 1990, 1991; Burchell, 1991; Ding et al., 1991; Ben-Arie et al., 1993). Several biotransformation enzymes have been identified in insects. These enzymes have features

780 Molecular Basis of Pheromone Detection in Insects

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suggesting they serve a dual function of attacking both xenobiotics and odor molecules. One of these, MsexGSTolf, is an antennal-specific GST in M. sexta capable of transforming aldehyde odorants (accession no. AF133268) (Rogers et al., 1999). GSTs often function as detoxification enzymes, complexing xenobiotics to endogenous glutathione (a cofactor) and thereby rendering them harmless (Yu, 1983, 1984, 1989; Fournier et al., 1992; Snyder et al., 1995). Though present in male and female antennas, in male antenna MsexGSTolf is restricted to cells underlying the pheromone sensitive trichoid sensilla (Figure 5a); this restricted expression to pheromone sensilla suggests that MsexGSTolf plays a role in pheromone inactivation (Rogers et al., 1999). MsexGSTolf mRNA encodes a 219 amino acid protein which lacks an encoded leader sequence; the absence of a leader sequence suggests that GSTolf is not extracellular but rather is retained in the cytoplasm of the cells expressing it. Pheromone molecules entering these cells, perhaps carried by PBPs, would be complexed and inactivated by MsexGSTolf and thus prevented from later stimulating the neurons (Figures 2 and 8). MsexGSTolf shares significant similarity with an antennal GST from B. mori (accession no. AJ006502) (Krieger and Breer, 1998); MsexGSTolf and BmorGSTolf are distinct from other insect GSTs and thus may define a unique class of multifunctional biotransformation enzymes (Rogers et al., 1999). An antennal-specific cytochrome P-450 enzyme has been identified in the moth M. brassicae; like MsexGSTolf, this biotransformation enzyme was also observed to differentially associate with pheromone sensitive trichoid sensilla (accession no. AY063500) (Maı¨be`che-Coisne´ et al., 2002). Male antennal specific and pheromone degrading P-450 activity has also been observed in the beetle Phyllopertha diversa; this activity was membrane-bound, required the cofactor NAD(P)H, and was sensitive to the cytochrome P-450 inhibitors proadifen and metyrapone (Wojtasek and Leal, 1999b). Again, these are presumably enzymes strategically located for the inactivation of xenobiotics, but may have become secondarily selected for pheromone degradation. Discussion earlier in this section described noctuid ALDHs that require cofactors (Tasayco and Prestwich, 1990a, 1990b). This requirement for cofactors in enzymes such as GSTs, P-450s, and ALDHs could be difficult to satisfy in the extracellular space of the sensilla lumen and may therefore suggest a cytosolic location of these enzymes. A cytosolic location of an ODE may seem incompatible with its function, i.e., once an odor/pheromone passes out of the sensillum lumen and enters a

cytoplasm it would seem effectively inactivated and out of reach from the perspective of the ORs. Perhaps this is not so. The restricted association of MsexGSTolf or M. brassicae P-450 with pheromone sensilla may be the consequence of strong evolutionary selection acting on pheromone systems to maintain low signal backgrounds. Cytosolic signal degradation would ensure that pheromone molecules entering the support cells could not later diffuse back into the sensillum lymph or activate ORs located in the nearby cell bodies of the olfactory neurons, actions that might disrupt reproductive behavior. 3.15.4.3.4. Category 4: cuticular ODEs – surface catabolism of background noise Airborne pheromone and other odors are hydrophobic and tend to adsorb onto the waxy surface of the insect cuticle. Body surfaces thus can collect odors and become sources of background noise if these odors are later released. Degradation of these surface-bound odor molecules might significantly reduce such signal noise. Pheromone degrading esterase activity associates with the scales covering the outer surface of the silk moth A. polyphemus (Vogt and Riddiford, 1986b). Isolated scales were loaded into small cartridges through which volatilized radioactive pheromone was blown ([3H]6E,11Z-HDA); alcohol metabolites were subsequently extracted indicating that the scales degraded adsorbed pheromone. Isolated scales were suspended in detergent solution containing either a-naphthyl acetate or [3H]-pheromone; both compounds were rapidly degraded. The degradative activity was apparently cross-linked to the scale cuticle as removal of the scales from the incubation solution reversibly removed the enzymatic activity as well. Boiling scales had only a partial effect on diminishing enzyme activity, suggesting that the presumed enzyme might be structurally stabilized by cross-linking to the cuticle. Activity was inhibited by trifluoroketone 1,1,1-trifluoro-2-tetradecanone and O-ethyl S-phenyl phosphoramidothiolate (EPPAT), the same compounds that inhibited the pheromone degrading of ApolSE (Vogt et al., 1985). Scales isolated from different body surfaces of A. polyphemus had somewhat different activities, and scales taken from other Lepidoptera had little or no activity compared to A. polyphemus, suggesting the activity was species specific for the A. polyphemus pheromone. These data suggested that the activity represented an essentially solid state and fairly indestructible enzyme selected to degrade sex pheromone adsorbed to the body surface, and that the A. polyphemus activity was

Molecular Basis of Pheromone Detection in Insects

tuned to the A. polyphemus pheromone (assuming the other species tested had their own scale esterases). This enzyme may function to reduce the noise that adsorbed pheromone might create if later released at an inappropriate moment. Surface catabolism of pheromones has been observed anecdotally but rarely studied with passion. Several studies have shown that homogenates or sonicates of general body parts of moths will degrade pheromone (Kasang, 1971; Mayer, 1975; Ferkovich et al., 1982). Kasang (1971) suggesting that this activity was due to surface enzymes acting to prevent adsorbed pheromone serving as a false signal source. Ferkovich et al. (1982) suggested that activity associated with legs of T. ni might serve to clean the antennas of residual pheromone by using the legs as antennal combs. Catabolism of a beetle pheromone, for example, has been observed by placing beetles elytra vials containing volatilized radioactive pheromone. Such enzymes should be of interest as solid state enzymes in applications of nonsolution chemistry or air filtration/detoxification. But they should also be interesting as examples of bizarre adaptive biochemistry underlying pheromone based behavior.

p0430

3.15.4.3.5. Inhibition of ODEs to inhibit pheromone behavior If ODEs can be differentially inhibited, then pheromone blend ratios might be altered within the sensillum and the pheromone thus misperceived and rendered unattractive. Several studies have used volatile trifluoroketones (TFKs) to probe the physiological and behavioral responses to lepidopteran pheromones; a TFK was shown to be a potent inhibitor of ApolSE (Vogt et al., 1985). These studies have produced variable results. A TFK applied directly to the antennas of the pyralid moth O. nubilalis partially inhibited whole antennal esterase activity but had no significant effect on male attraction to a pheromone source in a wind tunnel (Klun et al., 1991). However, attractant behavior was clearly inhibited in wind tunnel assays following pretreatment of the processionary moth (Thaumetopoea pityocampa) to volatilized TFKs (Quero et al., 1995), direct application of TFKs to the antennas of the noctuid moths Spodoptera littoralis and Sesamia nonagrioides (Bau et al., 1999), or copresentation with pheromone on a lure for the noctuid moth M. brassicae (Renou et al., 1999). The effectiveness of TFKs seems to relate to their structural similarity to the pheromone (Renou and Guerrero, 2000); copresentation of TFK and pheromone in field studies significantly reduced the attraction to traps for S. nonagrioides (Riba et al., 2001). The exact mode of action of TFKs is not clear; while they can clearly

781

inhibit esterase activities (Vogt et al., 1985; Vogt and Riddiford, 1986b), they may also interact with ORs (Klun et al., 1991; Pophof 1998; Pophof et al., 2000), and OBPs (untested). 3.15.4.4. Sensory Neuron Membrane Proteins

Sensory neuron membrane proteins (SNMPs) are antennal-specific proteins of Lepidoptera expressed in olfactory neurons and located in the receptive membranes of the dendrites (Figures 5f and 6). SNMPs belong to a larger gene family (CD36) that includes identified genes in other insects and other phyla (see below). SNMP (SNMP1) was first identified in the moth A. polyphemus (Rogers et al., 1997), and subsequently in the moths B. mori, H. virescens, and M. sexta (Rogers et al., 2001b); a second SNMP (SNMP2) was also identified in M. sexta (Robertson et al., 1999; Rogers et al., 2001b). These proteins express in both male and female antennas, although they are more abundant in male antennas, at least in A. polyphemus and M. sexta (not determined in B. mori or H. virescens). SNMPs are about 520 amino acids long; ApolSNMP1 is expressed as a 59 kDa peptide that is apparently posttranslationally modified to 69 kDa (Rogers et al., 1997, 2001b). The proteins are thought to contain two presumptive transmembrane domains located near the C- and N-terminals and a single large extracellular loop which contains several N-glycosylation groups and presumed disulfide bridges (Rogers et al., 1997, 2001b; Rasmussen et al., 1998; Tabuchi et al., 2000). The amino acid sequence identities between each SNMP1 range from 67% to 73%, and the identities between MsexSNMP2 and any of the SNMP1s are about 25%. Both SNMP1 and SNMP2 were recently identified in M. brassicae (MbraSNMP1 accession no. AF462066); MbraSNMP2 and Msex SNMP2 are about 65% identical (Jacquin-Joly, personal communication). SNMP expression suggests these proteins play a central role in odor detection. ApolSNMP1, MsexSNMP1, MsexSNMP2, and BmorSNMP1 are all known to be antennal specific (Rogers et al., 1997, 2001b). ApolSNMP1, MsexSNMP1, MsexSNMP2, and HvirSNMP1 are all known to express in olfactory neurons, and ApolSNMP1 protein has been visualized in the receptor membranes of these neurons (Rogers et al., 1997, 2001a, 2001b; Krieger et al., 2002) (Figure 5). ApolSNMP1 and MsexSNMP1 have been shown to be expressed late in adult development and into adult life, after morphogenesis has been completed but coincident with the expression of M. sexta OBPs and AOX and the onset of olfactory function (Vogt et al., 1993;

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782 Molecular Basis of Pheromone Detection in Insects

Figure 6 SNMP immunolabeling of electron micrograph sections of pheromone-sensitive trichoid sensilla. SNMP antibodies were visualized using secondary antibody conjugated to 10 nm colloidal gold particles. (a) and (b) include sensillum cuticle; (c) and (d) show only dendrites. Sensilla contained two neurons; one neuron consistently showed significantly greater labeling. Scale bars: (a) 1.25 mm, (b) 2.5 mm (c, d) 0.5 mm. (Modified from Rogers, M.E., Steinbrecht, R.A., Vogt, R.G., 2001a. Expression of SNMP-1 in olfactory neurons and sensilla of male and female antennas of the silkmoth Antheraea polyphemus. Cell Tissue Res. 303, 433–446.)

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Schweitzer et al., 1976; Rogers et al., 1997, 2001b). HvirSNMP1 expression coincides with the expression of OR genes in adult male H. virescens (Krieger et al., 2002). The unique and abundant association of SNMPs with the receptor membrane of olfactory neurons at a time when these neurons are capable of detecting and responding to odors suggests that SNMP are involved in odor detection. What do SNMPs do? The identification of ApolSNMP1 followed photoaffinity labeling studies that tentatively identified a 69 kDa protein as a pheromone receptor (Vogt et al., 1988) (Figure 5f); however, a role as pheromone receptor seems highly unlikely because SNMPs appear to associate with most olfactory neurons, and are neither 7-transmembrane domain receptors nor show the diversity expected for ORs. SNMPs certainly show no similarity to the presumed ORs identified in D. melangaster, A. gambiae and H. virescens (Clyne et al., 1999; Vosshall et al., 1999; Hill et al., 2002; Krieger et al., 2002). If SNMPs are not ORs, what are they?

SNMPs are homologous to a family of receptor proteins characterized by the human fatty acid transporter (FAT) CD36 (Oquendo et al., 1989; Greenwalt et al., 1992; Abumrad et al., 1993; see also Rogers et al., 2001a) (Figure 7). Among vertebrates, the CD36 family includes proteins known as scavenger receptors including LIMP II (Vega et al., 1991; Sandoval et al., 2000; Tabuchi et al., 2000), SRB1 (Acton et al., 1994, 1996; Krieger, 1999), and CLA 1 (Calvo and Vega, 1993). Members of this family have also been characterized in the insect D. melanogaster (Hart and Wilcox, 1993; Franc et al., 1999; Kiefer et al., 2002) and the slime mold Dictyostelium discoideum (Karakesisoglou et al., 1999; Janssen et al., 2001; Janssen and Schleicher, 2001). The function of the vertebrate CD36 proteins is becoming clear. In mammals, these proteins are reported to be receptors for both high and low density lipoproteins (HDLs and LDLs), transporters of cholesterol and phospholipids, thrombospondin receptors, and cell recognition receptors for phago-

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Figure 7 A neighbor-joining tree including lepidopteran SNMPs and their homolog in the Drosophila melanogaster and Anopheles gambiae genomes, as well as representative homolog from vertebrates (mammals) and a nematode (Caenorhabditis elegans). Box surrounds a similarity group that includes the olfactory specific lepidopteran SNMPs; Dmel-CG7000 is also known to be antennal specific (Vosshall, personal communication). Branches are collapsed to 50% bootstrap value or greater, based on 1000 replicates.

cytosis and recognition of Plasmodium (malaria) infected erythrocytes. Its role in malaria response was the first property ascribed to CD36 (Oquendo et al., 1989); CD36 is expressed in endothelial cells of blood vessels and is responsible for cytoadherence of infected erythrocytes to these endothelial cells (Gamain et al., 2002; Udomsangpetch et al., 2002). But the role of these proteins most studied is their involvement in heart disease, and specifically the formation of vascular plaques in atherosclerosis. Plaque formation involves lipid/cholesterol uptake by macrophages; this uptake converts the macrophages to foam cells; both cell phenotypes secrete growth factors which stimulate the formation of plaques. CD36 and SRB1 are expressed in these macrophages and are responsible for the binding of HDLs and LDLs to the macrophage, as well as the transport of cholesterols (components of the HDLs and LDLs) both into and out of the macrophage, either by forming channels (Calvo et al., 1998; Gu et al., 1998; Coburn et al., 2001; Reaven et al., 2001; Frank et al., 2002; Ibrahimi and Abumrad, 2002; Liu and Krieger, 2002; Miyazaki et al., 2002; Moore et al., 2002; Podrez et al., 2002a, 2002b; Tsukamoto et al., 2002; Kuniyasu et al., 2002) or by endocytosis (e.g., Kuniyasu et al., 2003). Similar activities of these proteins are now being examined in other contexts, including the mammalian nervous system with possible involvement in the mediation of

microglial and macrophage responses to b-amyloid, playing roles in proinflammatory events associated with Alzheimer’s disease (e.g., Panzenboeck et al., 2002; El Khoury et al., 2003; Srivastava, 2003). So, members of the vertebrate CD36 family can bind protein–lipid complexes and transport lipids across the cell membrane. These proteins have other less studied functions and activities as well. CD36 is involved in cytoadhesion and phagocytosis of rod outer segments in the eye (Ryeom et al., 1996). LIMP II has some as yet unknown function in lysosomes (Kuronita et al., 2002). SRB1 has recently been implicated as a receptor for hepatitis C virus (Scarselli et al., 2002). About 14 SNMP/CD36 homologs are present in the genomes of both Drosophila melanogaster and A. gambiae (Figure 7). Three of the D. melanogaster proteins have been characterized: emp (Hart and Wilcox, 1993), Croquemort (Franc et al., 1996, 1999; Lee and Baehrecke, 2001), and the product of ninaD (CG31783) (Kiefer et al., 2002). Emp (epithelial membrane protein) expresses in ectodermally derived tissues in the embryo and larva, including epithelial cells of wing imaginal discs; the authors suggested emp might have a role in development, but little more is known about this protein (Hart and Wilcox, 1993). Croquemort (Crq, catcher of death) expresses in circulating macrophages which engulf apoptotic cells by phagocytosis during

784 Molecular Basis of Pheromone Detection in Insects

nonautophagic cell death; Crq serves, perhaps along with other receptors, to mediate this process (Franc et al., 1996, 1999; Lee and Baehrecke, 2001). Although the precise role of Crq in this process is not known, the similarity between this activity and the phagocytosis activity of CD36 in mammals suggests that functions of this gene family are conserved between these two distant phyla (Ryeom et al., 1996; Franc et al., 1999). ninaD is a blind mutant that is deficient in the dietary uptake of the visual pigment precursor carotenoids; the ninaD mutation is in the gene CG31783, a CD36 homolog; P-element mediated transformation with wild-type CG31783 rescues ninaD mutants (Kiefer et al., 2002). NinaD expression was detected in embryonic midgut primordia, in mesodermal tissue giving rise to hemocytes and macrophage, and in hemocytes. The authors suggest that NinaD functions in the uptake of carotenoids, in a manner similar to the uptake of cholesterol and phospholipids by mammalian CD36 and SRB1 proteins (Frank et al., 2002; Liu and Krieger, 2002). The lepidopteran SNMPs are the only CD36 homologs clearly identified so far in neurons. Given what is known about the anatomy and physiology

of olfactory sensilla (Steinbrecht, 1999), it is difficult to translate the specific described activities of the mammalian and dipteran members of the CD36 gene family to olfactory function. However, in a general sense, members of this gene family function as receptors in cell–cell interaction, as receptors that bind protein–lipid complexes, and as transporters of small hydrophobic molecules such as cholesterol and probably carotenoids. Translating these activities into olfactory activities to formulate hypotheses of SNMP function is feasible (Figure 8). One Drosophila and two A. gambiae proteins share notable sequence similarity with the lepidopteran SNMPs (Figure 6), and it may prove that studies of these dipteran proteins, and especially CG7000 of Drosophila, may prove fruitful in elucidating SNMP function. 3.15.4.5. Perireceptor Models

Figure 8 illustrates several versions of the perireception scheme presented earlier (Figure 2) and redrawn in Figure 8a. In these schemes, OBPs transport odors to receptors and ODEs degrade odors where and when they have the opportunity. A major concern regarding OBP function is the

Figure 8 Models of ODE, OBP, and SNMP activities. (a) Depicts the general model for OBP and ODE activities proposed by Vogt et al. (1985); support cell activity is based on Rogers et al. (1999). Odor molecules are transported to ORs by OBPs and degraded by ODEs; odor molecules entering support cells are inactivated by dual function enzymes such as GSTs which also attack xenobiotics. (b) and (c) suggest alternative schemes for OBP activity (from Vogt et al., 1999), where the ORs are stimulated by a stable odor–OBP complex (b), or by free odor after the odor–OBP complex is destabilized by some other process such as interaction with SNMP (c) or cell membrane (Wojtasek and Leal, 1999a). (d–g) Several possible functions for SNMP (from Rogers et al., 2001a). SNMP may act as a novel odor receptor for either free odor (d-i) or odor–OBP complex (d-ii); or may destabilize the odor–OBP complex freeing odor to interact directly with ORs (e). Alternatively, SNMP may complex with ORs or cytosolic proteins to contribute in some manner to odor reception (f) or serve as an internalization process bringing odor or odor–OBP complexes into the cell (g). Further details on these models can be read in Rogers et al. (2001a, 2001b), and are discussed in the text.

Molecular Basis of Pheromone Detection in Insects

reversibility of binding within a physiologically relevant period of time (10–500 ms); thus, receptor activation is represented as either a simple transfer of pheromone from OBP to OR (Figure 8a), involving the direct interaction of a odor–OBP complex (Figure 8b) or following destabilization of the odor– OBP complex mediated through interactions with the membrane or other membrane proteins such as SNMP or some OR partner such as Dor83b (Figure 8c). Some of these alternatives have been considered in a mathematical model developed by Kaissling (e.g., Kaissling, 2001). Several schemes for SNMP action are shown in Figure 8d–g. Figure 8d considers the unlikely scheme that SNMP functions as an odor receptor, binding either (1) free odor or (2) the odor–OBP complex, while Figure 8e considers SNMP as a destabilizer of the odor–OBP complex. Perhaps more interesting is the scheme considered in Figure 8f where SNMP interacts with the OR or with intracelluar proteins (perhaps a guanylate cyclase, as described by Simpson et al. (1999)), or in some context similar to heterodimer receptor interactions observed in other systems (Vosshall and Keller, 2003). Finally, Figure 8g considers that SNMP might function to internalize odors or odor– OBP complexes, perhaps functioning as part of a signal termination pathway that removes odor accumulating near the membranes and thus maintains a concentration gradient for odors towards the neurons. This would be similar to the carotenoid transport function of NinaD in Drosophila, or the cholesterol transport function CD36 and SRB1 in mammals (Frank et al., 2002; Kiefer et al., 2002; Liu and Krieger, 2002; Kuniyasu et al., 2003). Not shown is the possibility that SNMP1 and SNMP2 serve as phenotype determinants of the respective neurons, providing a communication link between the sensilla neurons (cell–cell interaction) that sustains the identity of the respective neurons. These schemes are described in greater detail in Rogers et al. (2001a).

3.15.5. Conclusions 3.15.5.1. Does the Detection of Pheromones Differ from the Detection of Other Odor Molecules?

For many years, detection of pheromones was viewed as a unique system, distinct from the detection of odor molecules. However, pheromones are odor molecules, and the detection of pheromone at the level of the sensillum (i.e., periphery) appears to be an adaptationally selected version of the detec-

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tion of odors in general. Similarly, the central processing of pheromone and other odor signals has been argued to represent variations on a common theme (Christensen and Hildebrand, 2002). Nearly all of the elements illustrated in Figure 2 have homologs identified in both pheromonal and nonpheromonal systems. OBPs have been identified in orthopteroid and holometabolous/hemipteran lineages suggesting they are present at least throughout the neopterous insects, though their rare encounter in orthopteroid insects suggests they may have expanded considerably in the holometabolous lineages. One lineage of OBP, the PBPs and GOBPs, appears to be specific to the Lepidoptera and represents by far the most abundant OBPs present in lepidopteran antennas despite evidence that there are many more OBP genes expressed in the antennas (Vogt et al., 1991a; Robertson et al., 1999). Pheromone-specific lineages of OBP genes may also exist in other species, as suggested by distinct PBP/OBP lineages in beetle (Figure 3). ODEs have also been identified associating with both pheromone and nonpheromone sensilla, though there is clearly an enhancement in these enzymes in pheromone sensilla, at least in the Lepidoptera. ORs so far identified are yet to be associated with pheromone ligands, but the transduction pathway illustrated in Figure 2c has been shown to support both pheromone and nonpheromonal transduction in insects spanning the neopteran lineages. Pheromone ORs may yet prove to be derived (distinct in sequence and structure) from other ORs, but they are strongly predicted to be GPCRs activating and to be activated by similar transduction and perireceptor pathways as the nonpheromone ORs that have been characterized in Drosophila. It therefore seems appropriate now to do away with the old distinction of specialist (ligand equals pheromone) and generalist (ligand equals plant volatile) olfactory receptors (i.e., sensilla and OSNs) and instead focus on the commonalities of these systems and how the various pathways have become selected to detect and pass on sensory information in the context of life history needs of the species. 3.15.5.2. Are OBPs Necessary for OR Activation?

The behavioral response to ethanol of the Drosophila lush mutant has been considered as a strong support for a critical role for OBPs in odor detection (Kim et al., 1998; Kim and Smith, 2001). Assuming that Lush functions to transport ethanol to ORs, these experiments suggest OBPs are important in odor detection, but do not distinguish between (1) transport and (2) OBP–OR interaction. Certain-

786 Molecular Basis of Pheromone Detection in Insects

ly, odor detection will be influenced by the effectiveness of any system delivering odor molecules to the ORs, and the diversity and selectivity of OBPs seems to suit them to this role. However, several studies have suggested, or at least considered the possibility, that the odor–OBP complex functions as an intact unit in stimulating ORs (Prestwich et al., 1995; Ziegelberger, 1995; Plettner et al., 2000; Kowcun et al., 2001). Odor–OBP–OR interaction implies a high degree of stability in the odor– OBP complex, a view supported by several recent studies (Wojtasek and Leal, 1999a; Damberger et al., 2000; Sandler et al., 2000; Leal, 2003). Lazar et al. (2002) have gone so far as to suggest that such stability of the odor–OBP complex argues that OBPs are not deliverers of ligand (i.e., transporters), but rather are sequesters of ligand (inactivators), although this conjecture was based on a rather extreme extrapolation from elephant OBPs to moth OBPs, systems which share no homology in gene or species (beyond a divergence in their phylogenetic lineages perhaps at least 1 billion years ago) (Fortey et al., 1997; Erwin and Davidson, 2002). Several studies suggest OBPs may not be directly required for OR activation. In one study, olfactory neurons grown in primary culture responded to pheromone in the absence of applied PBP (Stengl et al., 1992). These cultures were established by dissociating cells of an early developmental stage of adult male antennas of the moth M. sexta. The authors questioned the necessity of PBPs in pheromone transduction, but also observed that PBP secreting cells may have been present and that PBP thus may have been present in the culture. However, under somewhat analogous conditions, Drosophila OR Dor43a was expressed in frog oocytes and stimulated with applied odors (Wetzel et al., 2001). These experiments were done without any applied OBP, and in the absence of any cells that could express an OBP, suggesting that the odor–OR interaction did not require a specific interaction with OBP. In a third study, the Drosophila OR Dor43a was misexpressed in olfactory sensilla lacking their endogenous ORs Dor22a/Dor22b (Dobritsa et al., 2003). Neurons of Dor22a/ Dor22b sensilla responded to Dor47a odor molecules, suggesting that the endogenous Dor22a/ Dor22b OBP(s) was capable of transporting the Dor47a ligand. If Dor22a/Dor22b and Dor47a sensilla express different OBPs, which is not currently known, then presumably if the Dor22a/Dor22b OBP was transporting the Dor47a ligand, it was doing so without a high degree of specificity. Also, if OBP–OR interactions do occur, this experiment suggests they do not occur with a high degree of

specificity. However, as there are as yet no correlations between the coexpression of ORs and OBPs, it remains possible that the Dor22a/Dor22b and Dor47a OBPs are the same, and that these assumptions are not supported. 3.15.5.3. Regulation of Pheromone Detection by Hormones, and the Regulation of Hormones by Pheromones

This subject falls into two broad areas: the hormonal regulation of olfactory development and gene expression, and the hormonal regulation of olfactory activity. Neither area has been heavily explored in insect olfactory systems, but sufficient work has been carried out to suggest there is much here to be studied. One can expect that development of the embryonic/larval olfactory system, modifications to the olfactory systems of hemimetabolous insects in the ultimate molt from juvenile to feeding adult, and metamorphosis of the olfactory systems of holometabolous insects offer rich landscapes to explore in an otherwise well-established background of endocrine studies (Truman and Riddiford, 2002). It is known that expression of the moth M. sexta PBP1, GOBP1, and GOBP2 are induced by declining levels of ecdysteroids late in adult development (Vogt et al., 1993) and that MsexGOBP2 expression is turned off and on again during a larval molt as if it were being regulated by ecdysteroids (Vogt et al., 2002). And it is known that several other olfactory genes including MsexAOX, MsexGSTolf, and MsexSNMP1 turn on coincident with the M. sexta OBPs, suggesting they too may be under ecdysteroid regulation (Rybczynski et al., 1990; Rogers et al., 1999, 2001b). We also know that eversion of the M. sexta antennal imaginal disc during the fifth larval instar is ecdysteroid sensitive (Fernandez and Vogt, unpublished data) as is the proliferation of antennal sensory cells shortly after pupation and early during adult development (M.D. Franco and R.G. Vogt, unpublished data). These studies verify that the ecdysteroid (and juvenile hormone) regulation of olfactory development offer significant research opportunities. The hormonal regulation of olfactory activity is largely unexplored territory. Insects display strong circadian behavioral shifts; moths, for example, have long been known to display sex pheromonerelated behaviors at night (Fabre, 1916; Rau and Rau, 1929) (see Chapter 3.11). While it is well established in Lepidoptera that pheromone release and pheromone associated behaviors are strongly regulated in a circadian fashion, there are no reports of circadian changes in the OSN response to sex pheromones. There is, however, a study in Drosophila demonstrating circadian changes in the antennal

Molecular Basis of Pheromone Detection in Insects

response to several odor molecules, though the function for this is only speculated to relate to feeding ecology (Krishnan et al., 1999; also discussed by Zwiebel, 2003). There is also recent data from A. gambiae about changes in the expression of a number of olfactory-related gene products following a blood meal (Justice et al., 2003). While these responses may be under local control, they may also be subject to endocrine pathways. Interestingly, the antenna, like most insect appendages, has a pulsatile organ at its base to move hemolymph through the antenna (Pass, 2000). In some insects at least, this also serves as neurohemal organ; hormones released from neurosecretory cells terminating in this structure are sent directly to the antenna, presumably to act in some as yet unknown fashion on antennal cells (Beattie, 1976; Pass et al., 1988a, 1988b; Pass, 2000). Hormones can certainly come from other sources in the insect, but this proximity of neurosecretory cells relative to the antenna seems a strong, if indirect, suggestion that the olfactory activities are under hormonal regulation. Pheromones have been categorized as releasers, stimulating or modulating an immediate behavioral response, and as primers, stimulating or modulating a longer-lasting physiological state (Wyatt, 2003). The best-characterized releaser pheromones may be the attractant sex pheromones of Lepidoptera which modulate visually guided flight of male moths (Kennedy et al., 1980; David et al., 1983; Preiss and Kramer, 1983; Willis and Arbas, 1991; Vickers et al., 2001; Baker, 2002; review: Schneider, 1992). Primer pheromones regulate behavioral and developmental states in social insects (Vander Meer et al., 1998) Leptinotarsa decemlineata (see Chapter 3.13). Pheromones may serve both categories, such as a male pheromone of the Colorado potato beetle, which serves as a female attractant (releaser), but may also modulate male pheromone production through an endocrine pathway (primer) (Dickens et al., 2002). Activities of releaser pheromones are presumably mediated through the interaction of pheromone-sensitive neural pathways with other sensory pathways to modulate motor output (Olberg, 1983; Kanzaki and Shibuya, 1986; Olberg and Willis, 1990, Kanzaki et al., 1991; Vickers and Baker, 1994; Kanzaki and Mishima, 1996; Kanzaki, 1998; Mishima and Kanzaki, 1998, 1999; Lei et al., 2001). Activities of primer pheromones are presumably mediated through the endocrine system. Unfortunately, there is currently no established neural, molecular, or genetic pathway between specific olfactory sensilla/OSNs/pheromones and any endocrine pathway. Nevertheless, this is a relatively easy pathway upon

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which to speculate. Neuroendocrine cells are distributed throughout the insect brain (Na¨ ssel, 2002), and projection neurons from the olfactory lobes arborize in regions rich in these cells (Figure 1); certainly the opportunity for olfactory-based signaling to influence the release of neurosecretory hormones is considerable. Given the state of genetically driven cell markers (Wong et al., 2002), all that may be lacking is a well-established pheromone responsive OR and the knowledge of the specific hormone(s) that pheromone simulates in order to establish such a neural pathway. New technologies drive the acquisition of new knowledge. Publicly accessible computer databases, N-terminal sequencing, cDNA cloning, PCR, and structural and spectroscopic biochemistry have generated much of the information presented here. The last few years have seen Drosophila emerge as a major tool in olfactory research, both because of its manipulatable genetics and its recently sequenced genome. The coming years will no doubt see these technologies spread to other insect models and the applications of profound new technologies to present models (Kalidas and Smith, 2002; Mori, 2002; Tomita et al., 2003). Changing technologies not only provide changes in research approaches, but also change that laboratories and which intellectual assets are serving as the central players of the moment. But for all this, insect olfaction is an inherently biological phenomenon. The vast number of insect species with their divergent families of olfactory genes that serve to translate key environmental information supporting individual life histories and behavior makes insect olfaction a unique system in which to study the organization, actions, and evolution of genes and genetic pathways in an ecological context. At the conferences, it is the rule rather than the exception to hear cutting edge presentations on gene and protein structure and properties, on gene regulation, expression, and function, on olfactory neural circuitry, coding and behavior, on the most superb neural anatomy, on development at all levels, on the chemistry of pheromone production and the evolution of pheromone production and detection, on ecology of odor detection, on field application of pheromones and crop protection using pheromones, and recently, on remote sensing and military defense. Perhaps more than any other area of biology, insect olfaction is a system offering and generating enormous intellectual integration. For this aging scientist, studying insect olfaction has been highly stimulating, and I can only envy those younger scientists now embarking in this field.

788 Molecular Basis of Pheromone Detection in Insects

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3.16

Endocrinology of Crustacea and Chelicerata

E S Chang, University of California, Bodega Bay, CA, USA W R Kaufman, University of Alberta, Edmonton, AB, Canada ß 2005, Elsevier BV. All Rights Reserved.

3.16.1. General Introduction 3.16.2. Crustacea 3.16.2.1. Introduction 3.16.2.2. Molting and Metabolism 3.16.2.3. Pigmentary Hormones 3.16.2.4. Reproduction 3.16.2.5. Limb Autotomy and Regeneration 3.16.2.6. Conclusions 3.16.3. Chelicerata 3.16.3.1. Introduction 3.16.3.2. The Neuroendocrine Systems of the Chelicerata 3.16.3.3. Molting and Metamorphosis 3.16.3.4. Reproduction 3.16.3.5. Salivary Gland Development and Degeneration in Female Ixodid Ticks 3.16.3.6. A Role for 20-Hydroxyecdysone as a Reproductive Pheromone in Ixodid Ticks 3.16.3.7. The Role of ‘‘Mating Factors’’ in Tick Reproduction 3.16.3.8. Conclusions

3.16.1. General Introduction The Arthropoda comprise four subphyla (Mitchell et al., 1988): Trilobita (about 4000 extinct species), Crustacea (about 40 000 species), Uniramia (about 1 million species, most of them insects), and Chelicerata (about 65 000 extant species). Although the focus of this series is on the insects, the editors of this volume decided that any thorough treatment of insect endocrinology should include a comparative discussion of the hormones of the other major extant arthropod groups. Some of the hormones described here are ubiquitous among the Arthropoda (e.g., the ecdysteroids). There are several other hormones, however, that appear to be unique in these groups. Of course these groups cannot be treated as comprehensively in a single chapter as can the insects in an entire volume.

3.16.2. Crustacea 3.16.2.1. Introduction

The major crustacean classes are the Branchiopoda (water fleas and brine shrimp), the Maxillopoda (copepods and barnacles), the Ostracoda, and the Malacostraca (amphipods, isopods, and decapods).

805 805 805 805 811 812 813 813 814 814 814 816 822 826 828 828 829

The decapods include the familiar crabs, crayfish, shrimp, and lobsters. Crustaceans are found in a wide variety of habitats that include marine, freshwater, and terrestrial environments and range from deep-sea thermal vents to freshwater pools in lightless caves. The adult head of crustaceans bears first and second antennae, mandibles, and first and second maxillae. Various types of appendages can be found on the thorax and abdomen. Although there are numerous orders of crustaceans, the greater part of the endocrinological work has been conducted on the Decapoda. 3.16.2.2. Molting and Metabolism

Arthropods must periodically shed their external, confining exoskeletons and take up air or water to expand their new and larger exoskeletons in order to grow in size. The problem of how to increase in body size is even more formidable for crustaceans due to their mineralized, relatively rigid exoskeletons. The molt cycle is defined as the interval between two successive molts. Drach (1939) divided the molt cycle into a series of stages associated with specific cuticular events from just after ecdysis (postmolt), through a lengthy intermolt period, to the period leading up to a subsequent ecdysis (premolt). These stages have been assigned a number of

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substages, depending upon the author and the species of crustacean (e.g., Aiken, 1973, for the lobster Homarus americanus). 3.16.2.2.1. Ecdysteroids Horn and his colleagues (Hampshire and Horn, 1966; Horn et al., 1966) isolated and determined the structure of 20hydroxyecdysone (20E) from the rock lobster Jasus lalandei. Although the hormone was originally called ‘‘crustecdysone’’ or ‘‘ecdysterone,’’ Horn’s group demonstrated the identity of 20E as the principal active form of the molting hormone from both insect and crustacean sources. In most species examined, ecdysone (E) is the prohormone secreted by the Y-organ and it is hydroxylated by target tissues to 20E (Chang and O’Connor, 1978). There are exceptions to this, however, as described below. Although 20E is the predominant molting hormone in all decapod species examined to date, other ecdysteroids (steroids structurally related to E that have molting hormone activity) have been characterized in hemolymph and tissues of various crustacean species. These include E; ponasterone A (25-deoxy-20hydroxyecdysone); inokosterone (25-deoxy-20,26dihydroxyecdysone); makisterone A (24-methyl20-hydroxyecdysone); 20,26-dihydroxyecdysone; and 2-deoxyecdysone (reviews: Spindler et al., 1980; Skinner, 1985a; Watson et al., 1989; Chang, 1993). Comparisons in the absolute concentrations of various ecdysteroids between species are difficult to make due to different detection methods and the lack of standardization in the molt stage of the donor animals. Classical morphological observations and endocrinological experiments (Gabe, 1953; Echalier, 1959) indicated that the molting gland in the crab Carcinus maenas was the thoracic Y-organ. Organ culture of the Y-organs from the crabs Cancer antennarius and Pachygrapsus crassipes (Chang and O’Connor, 1977) and from the crayfish Orconectes limosus (Keller and Schmid, 1979) resulted in the characterization of E as the primary secretory product of the Y-organ. Other ecdysteroids are secreted by the Y-organs in other crab species: 3-dehydroecdysone from Cancer antennarius (Spaziani et al., 1989) and 25-deoxyecdysone from Carcinus maenas (Lachaise et al., 1989; see review by Watson et al., 1989). It is not clear why different types of ecdysteroids are secreted by the Y-organs of various species. In almost all cases examined, exogenous ecdysteroids promote progression through the molt cycle (Skinner, 1985b). Associated biological activities attributed to exogenous ecdysteroids include apolysis (separation from the exoskeleton from the underlying

epidermis) (Webster, 1983), growth of regenerating limb buds (Rao et al., 1972; Hoarau, 1982), formation of gastroliths (McWhinnie et al., 1972), increased incorporation of labeled precursors into macromolecules (McWhinnie et al., 1972; Dall and Barclay, 1979), increased protein kinase activity (Christ and Sedlmeier, 1987), premolt behavior (Borowsky, 1980), and alterations in hemolymph osmolarity (Charmantier and Trilles, 1976). Like vertebrate steroid hormones, ecdysteroids recognize target tissues by binding with nuclear receptors. The ecdysteroid receptor has been isolated and characterized from the fiddler crab Uca pugilator (Durica and Hopkins, 1996; Chung et al., 1998). It has been sequenced and has homologies with insect ecdysteroid receptors (see Chapter 3.5). Transcripts for the crab gene were isolated from limb buds and developing ovaries (Durica et al., 2002). 3.16.2.2.2. Molt-inhibiting hormone Hemolymph ecdysteroid concentration fluctuates dramatically during the molt cycle (e.g., from 350 ng ml
1 in premolt lobster) (Chang and Bruce, 1980) and these changes mediate the various biochemical and physiological processes that occur during the cycle. The rate of synthesis and/or secretion of E by the Y-organ varies during the molt cycle and partially explains these hemolymph fluctuations in ecdysteroid titer. Just prior to the substage of premolt in which the highest concentration of ecdysteroids was observed in the hemolymph, explanted Y-organs were found to secrete the greatest amount of E (Chang and O’Connor, 1978). Low hemolymph concentrations were correlated with low secretory rates. Removal of both stalked eyes of U. pugilator resulted in a shortening of the molt interval (Zeleny, 1905). This observation lead to the postulation of an endocrine factor present in the eyestalks that normally inhibits molting – a molt-inhibiting hormone (MIH). Detailed microscopical examinations resulted in the description of a neurohemal organ in the eyestalk of several decapod crustaceans (Bliss and Welsh, 1952; Passano, 1953). This neurohemal organ is called the sinus gland and serves as a storage site for neurosecretory products. It consists of the enlarged endings of a group of neurosecretory neurons collectively called the X-organ (Hanstro¨m, 1939). The shortened molt interval observed in eyestalkablated decapods is likely due to a rapid elevation in the concentration of circulating ecdysteroids, which is a result of X-organ/sinus gland removal (Chang et al., 1976; Chang, 1993). Conversely, the

Endocrinology of Crustacea and Chelicerata

demonstration of decreased ecdysteroid titers in eyestalk-ablated crabs that have been injected with eyestalk extracts (Hopkins, 1982; Keller and O’Connor, 1982) lends additional support to the paradigm of the MIH–Y-organ molt-controlling axis. Further evidence of this hypothesis was provided by Gersch et al. (1980), who demonstrated that if O. limosus were initially injected with sinus gland extracts, the subsequent culture of the crayfish Y-organs resulted in a decreased production of E compared to vehicle-injected animals. In vitro secretion of E by crab Y-organs could be inhibited when cultured with either conditioned medium that had previously been incubated with explanted sinus glands or by the addition of eyestalk extracts (Soumoff and O’Connor, 1982; Mattson and Spaziani, 1985; Schoettker and Gist, 1990). Callinectes sapidus MIH has been cloned and the cDNA was used as a probe to show that the amount of MIH mRNA in the eyestalk neural ganglia was negatively correlated with the hemolymph concentration of ecdysteroids; for example, there were low levels of MIH mRNA in the eyestalk during premolt (Lee et al., 1998). Similarly, Nakatsuji et al. (2000) observed an increased content of MIH in the premolt X-organ/sinus gland in Procambarus clarkii. MIH from C. maenas was among the initial MIHs to be characterized (Webster, 1991) (Figure 1). It is a member of a novel neuropeptide family, representatives of which have so far been found only in arthropods (Edomi et al., 2002). This neuropeptide family regulates such diverse functions as molting, reproduction, and metabolism (Chang, 1993; De Kleijn and Van Herp, 1995; Webster, 1998; Watson et al., 2001; Bo¨ cking et al., 2002). MIH binds to hormone receptors on the membranes of Y-organ cells (Webster, 1993) and likely mediates its action via cyclic nucleotide second messengers (Saidi et al.,

807

1994) or through alterations in calcium ion fluxes and activity of protein kinase C that phosphorylates enzymes involved in steroidogenesis (Spaziani et al., 2001). (See Chapter 3.2 for the role of second messengers in the modulation of E synthesis in the insect prothoracic gland.) 3.16.2.2.3. Crustacean hyperglycemic hormone and related neuropeptides Crustacean hyperglycemic hormone (CHH), initially characterized from the crab C. maenas (Kegel et al., 1989; Weidemann et al., 1989), contains a high degree of homology with MIH (Figure 1). One important role of CHH is the maintenance of glucose homeostasis. Removal of the sinus glands from O. limosus, leaving the remainder of the eyestalk neural tissue and vision intact, resulted in a permanent decline in hemolymph glucose. The circadian rhythmicity of fluctuations in glucose concentration was attenuated (Hamann, 1974). Hyperglycemia is commonly observed under a variety of stressful conditions, and there is considerable indirect evidence to suggest that stress-induced hyperglycemia is caused by the release of CHH (Kleinholz and Keller, 1979; Keller and Sedlmeier, 1988; Webster, 1996). For example, under hypoxic conditions, glucose may be released by CHH from carbohydrate stores to serve as a substrate for glycolysis and lactate formation, as is typical for anaerobiosis of decapod crustaceans. After 4 h of emersion, which causes anoxia, the CHH concentration in lobsters rose from 55% identical) and an arabic numeral designates the individual gene (all italics) or message and protein (no italics) (Figure 1). Different P450 enzymes are generally products of different genes; they are not isozymes or isoforms. The identity (%) rules for family and subfamily designations are not strictly adhered to, but names once adopted are rarely changed. Initially, many insect P450s were arbitrarily lumped into the CYP6 and the CYP4 families even though they had less than 40% amino acid identity with CYP6A1 or with vertebrate CYP4 proteins. Naming genes in the lumper mode made the CYP6 and CYP4 families the largest ones in insects by a cascade effect. CYP6B1 is only 32% identical to CYP6A1 (Cohen et al., 1992), so placing it in the CYP6 family ‘‘forced’’ many subsequent sequences into that family even if they did not meet the 40% criterion. The splitter mode prevailed at the completion of genome projects, which led to a new proliferation of CYP families in insects, the CYP300 series. A termite P450 claimed the welcoming designation of CYP4U2 (GenBank AF046011). Gotoh (1993) has introduced a useful nomenclature of higher order than CYP families: the E (for eukaryotic type) and B (for bacterial type) ‘‘classes’’ and subclasses (I, II, III, etc.) that regroup CYP families on the basis of phylogeny. Nelson (1998) has similarly introduced the notion of ‘‘clans,’’ but the precise criteria for naming Gotoh’s ‘‘classes’’ and Nelson’s ‘‘clans’’ have not been defined. Alleles of a gene are named as subscripts v1, v2 (e.g., CYP6B1v2, Cohen et al., 1992). The human P450 polymorphisms are named according to a clear nomenclature. Pseudogenes are noted by the

4.1.1.4. Nomenclature

A nomenclature of P450 genes and proteins was introduced when only 65 sequences were known

Figure 1 Scheme of the P450 nomenclature.

4 Insect Cytochrome P450

suffix P. This suffix ought to be added to the closest paralog that is an active gene, e.g., CYP9E2 and CYP9E2P1 in Blattella germanica (Wen et al., 2001). However this is not always done, as the closest paralog is sometimes not easily recognized. In following the tradition that predates the CYP nomenclature, P450 enzymes can be named with a small suffix, such as P450cam, the camphor hydroxylase of Pseudomonas putida later named CYP101; P450BM3 the fatty acid hydroxylase of Bacillus megaterium (CYP102); or P450scc, the cholesterol side-chain cleavage enzyme (CYP11A1). In insects, few P450 enzymes have been named in this way. P450Lpr is the predominant P450 in the pyrethroid-resistant strain Learn-Pyr of the housefly, later identified as CYP6D1 (Tomita and Scott, 1995). P450hyd (Reed et al., 1994) is the P450 forming hydrocarbons in the housefly. P450MA is a P450 purified from the Munsyana strain of the German cockroach (Scharf et al., 1998). In the Drosophila gene nomenclature (Lindsley and Zimm, 1992), only the initial letter is capitalized, hence CYP6A1 in the housefly and Cyp6a2 in Drosophila. The CYP nomenclature of Nebert et al. was clearly designed to reflect the evolutionary relationships between the genes as evidenced by the degree of sequence identity of the proteins they encoded. As such it is a Darwinian nomenclature. P450 proteins have been categorized into classes (Ravichandran et al., 1993) that reflect the types of electron delivery to the active site. Class I proteins require both an FAD-flavoprotein reductase and a ferredoxintype protein, class II P450s require only an FAD and FMN diflavin reductase, class III enzymes are self-sufficient, and class IV P450s receive electrons directly from NADPH. The utility of this nonevolutionary classification is debatable.

4.1.2. Diversity and Evolution of Insect P450 Genes 4.1.2.1. Sequence Diversity

4.1.2.1.1. P450 sequences from classical cloning techniques The first insect P450s cloned and sequenced were CYP6A1 from Musca domestica in 1989 (Feyereisen et al., 1989), CYP4C1 from Blaberus discoidalis in 1991 (Bradfield et al., 1991), CYP6A2 and CYP4D1 from Drosophila (Gandhi et al., 1992; Waters et al., 1992) as well as CYP6B1 from Papilio polyxenes in 1992 (Cohen et al., 1992), and CYP4D2 from Drosophila in 1994 (Frolov and Alatortsev, 1994). The methods used in these early studies were screening of cDNA expression libraries with polyclonal (CYP6A1) or monoclonal

(CYP6A2) antibodies to (partially) purified P450 proteins of insecticide-resistant flies (CYP6A1, CYP6A2). For CYP6B1, microsequencing of Nterminal and internal sequences of a P450-sized band on a SDS-PAGE gel of P. polyxenes larval midgut proteins led to the design of degenerate oligonucleotide probes for RT-PCR on midgut poly(A)þ RNA. The cloned PCR product was used in turn as a probe to screen a xanthotoxin-induced midgut cDNA library. Significantly, two cDNAs representing alleles of the CYP6B1 gene were thus isolated. Classical molecular approaches of this type have continued to yield new P450 sequences (Tomita and Scott, 1995; Wang and Hobbs, 1995; Winter et al., 1999) and the initial P450 sequences have served as probes to isolate related sequences in the same species (Cohen and Feyereisen, 1995; Hung et al., 1995a, 1996; Maitra et al., 1996), or in phylogenetically close species (Li et al., 2000a, 2001). Interestingly, CYP9A1 of Heliothis virescens was cloned by screening an expression library with monoclonal antibodies that also served to clone CYP6A2 of Drosophila, despite the fact that the two sequences are only 32.4% identical (Rose et al., 1997). 4.1.2.1.2. Serendipity and P450 discovery In contrast to the targeted approaches, CYP4C1 was obtained in 1991 by differential screening of a cockroach fat body cDNA library (Bradfield et al., 1991). The probes consisted of cDNA obtained from fat bodies of hypertrehalosemic hormonetreated roaches, or decapitated controls. The cloning and identification of Drosophila Cyp4d1 and Cyp4d2 were equally serendipitous, as investigators were interested in transcripts in the prune region at the tip of the X chromosome (Gandhi et al., 1992; Frolov and Alatortsev, 1994). Drosophila Cyp18 was initially cloned in a screen for ecdysoneinducible genes (Hurban and Thummel, 1993) and subsequently obtained as full-length cDNA (Bassett et al., 1997). Cyp4e1 deserves a special historical mention. Its sequence was discovered independently in a 9 kb genomic sequence of the 44D region of Drosophila encompassing a cluster of cuticle protein genes (GenBank Acc. K00045) by T. Holton and D. Nero in 1991, each searching databases with P450 sequences (personal communications). This partial sequence of insect Cyp4e1 is thus the first (1983), though not annotated, record of an insect P450 in GenBank. 4.1.2.1.3. Exponential amplification of new P450 sequences by PCR By 1994, sufficient information about vertebrate and insect P450s allowed the

Insect Cytochrome P450 5

isolation of fragments of P450 cDNAs and genes by PCR methods with degenerate oligonucleotides corresponding to consensus sequences. A variety of approaches were taken, two of which being particularly successful: in the first, sequences in the I helix and surrounding the conserved cysteine (see Section 4.1.3.3.1) were used to isolate PCR products of about 450–500 bp that coded for about 130 amino acids or almost a third of the full-length P450. In the second, the sequence around the conserved cysteine was used to design a first primer, with oligo(dT) serving as anchor on the poly(A)þ message. Fragments of varying length were obtained by this 30 RACE strategy, the C-terminal 30–50 amino acids sequence of the P450 and a variable 30 UTR sequence (review: Snyder et al., 1996). The PCR approaches led to the description of 17 new CYP4 genes in A. albimanus (Scott et al., 1994); 5 CYP4 genes from Manduca sexta (Snyder et al., 1995); 8 CYP genes from H. armigera (Pittendrigh et al., 1997); 4 genes of the new CYP28 family from Drosophila species (Danielson et al., 1997); 14 P450 fragments from Ceratitis capitata (Danielson et al., 1999), 95 new sequences from 16 drosophilid species (Fogleman et al., 1998), etc. (Amichot et al., 1994; Stevens et al., 2000; Tares et al., 2000; Scharf et al., 2001; Wen et al., 2001; Ranson et al., 2002b). The PCR method has often been used as a first step in the isolation of full-length P450 clones in insects (Snyder et al., 1995; Danielson and Fogleman, 1997; Hung et al., 1997; Danielson et al., 1998, 1999; Guzov et al., 1998; Kasai et al., 1998a, 2000; Ranasinghe and Hobbs, 1998; Sutherland et al., 1998; Stevens et al., 2000; Wen et al., 2001; Wen and Scott, 2001b; Liu and Zhang, 2002) and ticks (Crampton et al., 1999; He et al., 2002). The large number of partial P450 sequences obtained by cloning and sequencing an even larger number of PCR products has led to two related problems. The first is that small sequence differences can be found between very closely related PCR products. Are these artifactual, or do they represent allelic variation? In the study on A. albimanus for instance, 64 clones were sequenced of which 47 encoded P450 fragments, describing 17 genes (Scott et al., 1994). In some clones, up to seven nucleotide differences from the closest sequence were seen. A few nucleotides may not just differentiate two alleles of the same gene, but may be sufficient to differentiate two genes, as the complete sequence of the Drosophila and A. gambiae genomes subsequently showed. This causes a nomenclature problem, when two distinct genes are prematurely described as allelic variants of a single gene. The second problem is that the P450 fragments obtained

by PCR, one third of the full sequence or less, make the calculation of percentage identity (the base of the nomenclature rules) difficult. In the A. albimanus study (Scott et al., 1994) this problem was resolved by establishing a function that derives identity over the full length P450 from the percentage identity of the PCR product. Further validations of this approach have been presented (Danielson et al., 1999; Fogleman and Danielson, 2000). Nonetheless, the practice of bestowing an official CYP designation to a short, partial P450 sequence should be abandoned. 4.1.2.1.4. Genome sequences: diversity revealed The D. melanogaster complete genome sequence was published in 2000 (Adams et al., 2000). Annotation of the P450 sequences (Tijet et al., 2001) was done by multiple Basic Local Alignment Search Tool (BLAST) searches, taking into account intronexon structure, pairwise and multiple alignments, EST sequences, and known features of wellcharacterized P450. This task gave 90 sequences, of which 83 appeared to code for potentially functional P450s. Seven sequences were either partial sequences or obvious pseudogenes. Forty genomic sequences were not represented by ESTs. With release 3 of the Drosophila genome (Celniker et al., 2002), the ‘‘official count’’ of potentially functional P450 genes is 85 with five pseudogenes. A similar annotation of the P450 genes from the complete A. gambiae genome published in 2002 (Holt et al., 2002) gave 111 genes (Ranson et al., 2002a) of which five are thought to represent pseudogenes. The need for ‘‘manual’’ annotation remains critical because of the error-prone gene- and transcriptcalling programs, as noted for P450s by Gotoh (1998). These difficulties remain a major challenge to the ‘‘completion’’ of a genome program (Misra et al., 2002). Potentially confusing is the proliferation of GenBank accessions for transcripts identified in silico, with only scant notice that they represent nothing but software-digested genomic sequences. Some of the sources of errors noted for the Drosophila annotation (Misra et al., 2002) were plainly evident with the A. gambiae genome sequence as well. They typically include the fusion of two neighboring P450 genes into one, or the truncation of a gene. Comparison of the intron/exon structure of closely related genes and alignments with EST sequences when these are available can facilitate P450 gene annotation in a majority of cases. Independent evidence, such as the cloning of full-length cDNAs, or functional expression is rarely available to resolve annotation problems, so that the complete description of the P450 gene complement of any

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species remains an ongoing task. A website presents information on the P450 sequences from completely sequenced genomes (D. melanogaster, A. gambiae) as well as links to other P450 websites. The complete sequence of an increasing number of insect genomes (Drosophila pseudoobscura, Apis mellifera, Bombyx mori, Tribolium castaneum) and the continuous addition of P450 sequences in EST projects from various species is redefining the approach to P450 research. 4.1.2.2. Genomic Organization: Clusters p0090

The presence of P450 genes in clusters was revealed in early studies with Drosophila and the housefly (Frolov and Alatortsev, 1994; Cohen and Feyereisen, 1995). Further evidence was obtained by in situ hybridization to polytene chromosomes of Drosophila (Dunkov et al., 1996) and ultimately by the analysis of P450 genes in the complete genome sequences of Drosophila (Tijet et al., 2001) and A. gambiae (Ranson et al., 2002a, 2002b). The largest Drosophila P450 cluster carries nine genes (eight CYP6A genes and CYP317A1) at 51D on the right arm of chromosome 2. Six of these genes (Cyp6a17 to Cyp6a21) are coordinately regulated during the circadian rhythm (Ueda et al., 2002). In A. gambiae, the largest cluster carries 14 P450 genes of the CYP6 family at 30A on the right arm of chromosome 3. A large cluster of CYP325 genes in A. gambiae contains 12 genes and 2 pseudogenes. In A. gambiae, only 22 of the 111 genes are present as singletons, with 16 clusters of 4 or more genes (Ranson et al., 2002a) (Figure 2). Gene clusters are thought to arise by sequential gene duplication events, the principal and initial mechanism of P450

Figure 2 Clusters of P450 genes and singletons (cluster size ¼ 1) genes in the Drosophila melanogaster (blue) and Anopheles gambiae (red) genomes. (Reproduced with permission from Ranson, H., Claudianos, C., Ortelli, F., Abgrall, C., Hemingway, J., et al., 2002a. Evolution of supergene families associated with insecticide resistance. Science 298, 179–181; ß AAAS.)

diversification. In Papilio glaucus, CYP6B4v2 and CYP6B5v1 are clustered within 10 kb of each other (Hung et al., 1996). They are 99.3% identical at the nucleotide level, with 98% identity of their single 732-bp intron and 95% identity over 616 bp of the promoter region. The proteins they encode differ by just one amino acid. CYP6B4 and CYP6B5 are thus recently duplicated genes that have not yet diverged substantially in sequence (Hung et al., 1996). Similar close pairs are found in Drosophila, e.g., Cyp12a4/Cyp12a5; Cyp9b1/Cyp9b2; Cyp28d1/Cyp28d2. Cyp12d1 and Cyp12d2 are 2 kb apart and differ by only three nucleotides leading to three changes at the amino acid level. In A. gambiae, CYP6AF1 and CYP6AF2 are 99.8% identical at the nucleotide level and differ by just one amino acid (Ranson et al., 2002a). CYP6D1 and CYP6D3 are clustered on chromosome 1 in the housefly (Kasai and Scott, 2001a). Although they are only 50% identical in their 500 nt UTR and their product 80% identical at the amino acid level, both genes are phenobarbital-inducible and are constitutively overexpressed in the LPR strain (Kasai and Scott, 2001b). In the housefly, a cluster of six CYP6 genes within 24 kb of each other on chromosome 5 shows evidence of both gene duplications and chromosomal inversions (Cohen and Feyereisen, 1995). Three genes are transcribed in one direction and the next three are transcribed in the opposite direction. The six genes have a short intron at the same position in the ETLR conserved region (see Section 4.1.3.3.1) as many CYP6 genes. The CYP6A6 gene located at one extremity of the cluster is in fact represented only by the second exon. The first exon was not found in 2 kb of DNA upstream of this exon boundary, although the intron length of the five other genes is only 57–125 bp. CYP6A6 may therefore be a pseudogene generated during a chromosomal inversion whose breakpoint was located in the intron. The other five genes of the cluster are transcribed, with CYP6A5 being predominantly expressed in larvae (Cohen and Feyereisen, 1995). Two recently duplicated genes may undergo gene conversion, if they do not diverge fast enough (Walsh, 1987). Gene conversion at the 50 end of the CYP6B8 and CYP6B28 genes of Helicoverpa zea (Li et al., 2002b) has been suggested by the lower level of nt differences in the first half of the first exon, as compared to the rest of the sequence. Exon specific deficit in variation among pairs of P450 is indicative of a gene conversion event (Matsunaga et al., 1990). Changes in regulatory patterns (induction, tissue specific expression) can be observed for recently duplicated genes (Li et al., 2002c) and may

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lead to their independent evolution. Extreme examples of gene conversion between duplicated P450 genes leading to their concerted evolution, of the kind seen with a-amylase genes (Hickey et al., 1991), has not been reported to date. Although events such as unequal crossing-over can lead to gene duplications, there are other mechanisms such as retrotransposition. Capture of a spliced mRNA by a retrotransposon can reintroduce an intronless sequence into the genome where it may evolve further or die as a pseudogene. This process generally occurs in germ cells, so it should be limited to genes that are expressed in those cells. The Drosophila Cyp4g1 gene at the tip of the X chromosome (1B3) may be a case in point. It lacks introns and its closest paralog, Cyp4g15, has five introns and is located at 10C2-3. Furthermore, Cyp4g1 is represented by the largest number of ESTs in Drosophila, and yet the apparent N-terminus of CYP4G1 is quite atypical of a microsomal P450 suggesting that an ancient 50 UTR is now part of the open reading frame. Further studies are needed to clarify the status of this gene. There is little evidence for alternative splicing of insect P450 transcripts as additional means of generating diversity. The Cyp4d1 gene seems to utilize two alternate first exons, and ESTs for each transcript type, CYP4D1 and CYP4D1alt, have been found. The first cDNA cloned (Gandhi et al., 1992) uses a first exon (exon 1 prime) closest to the rest of the gene, whereas several ESTs use a more distal first exon (exon 1) instead. The two predicted proteins differ only from the N-terminal to the beginning of the first SRS (see Section 4.1.3.3.1 and Figure 8). The consequences of this alternative splicing in terms of catalytic competence thus remain to be examined. 4.1.2.3. Genomic Variation: Alleles, Pseudogenes

Apparent allelic variants of cloned P450 cDNAs and genes were already described in the earliest studies of insect P450s (Cohen et al., 1992, 1994; Cohen and Feyereisen, 1995). Examples of this variation are most striking in A. gambiae, because the sequence released includes several ‘‘scaffolds,’’ which are not included in the ‘‘golden path’’ used for genome assembly. These scaffolds represent large tracts of heterozygosity (‘‘dual haplotype regions,’’ Holt et al., 2002), probably resulting from the mosaic nature (contributions from different A. gambiae cytotypes) of the strain that was sequenced. Interestingly, some of these scaffolds harbor P450 genes, even P450 gene clusters, and a comparison of the various haplotypes reveals not only considerable

differences, but also variations in the total number of P450 genes. In other words, P450 gene duplication events have occured independently in different mosquito cytotypes. A gene recently converted to a pseudogene in one population or strain, as a result of a debilitating mutation or transposable element insertion (see Chapter 4.12) may still be active in another population. The total number of P450 genes in a species is therefore a relative number that is genotype dependent. In Blattella germanica, three pseudogenes of CYP4C21 and two pseudogenes of CYP9E2 have been described (Wen et al., 2001). These pseudogenes are characterized by deletions and/or the presence of several stop codons in the open reading frame. CYP9E2P2, for instance, has just 10 nucleotide differences from CYP9E2, but two base deletions render this gene nonfunctional. The processed pseudogene CYP4W1P in the cattle tick has a 191 bp deletion, but only three other nt changes in the open reading frame (Crampton et al., 1999). Another tick pseudogene, CYP319A1P, contains two DNA insertions in the open reading frame and also appears to be of recent origin (He et al., 2002). The rate of gene duplication has been reported to be extremely high, making it an event almost as frequent as point mutations at the nucleotide level (Lynch and Conery, 2000). With a rate of 31 gene duplications per genome per million years in Drosophila, and a half life of less than 3 million years for a duplicated gene, one can estimate that there is a P450 gene duplication event on average every 5 million years. Drosophila is peculiar in its ability to delete genomic DNA at a high rate, and the proportion of pseudogenes in the genome is low – 1 pseudogene for 130 proteins as compared to 1 for 19 in Caenorhabditis elegans (Harrison et al., 2003). However the proportion of pseudogenes in the P450 family is high (five pseudogenes) in Drosophila, another indication of rapid turnover of P450 genes. 4.1.2.4. P450 Gene Orthologs

With so much gene duplication, are there still any P450 orthologs in related species? (orthology ¼ the ‘‘same’’ gene in different species that has only diverged as a result of speciation). The availability of two completely sequenced genomes made it possible to identify those members of the P450 family that were truly orthologous. This is not as easy as it seems, because in some borderline cases, two formally orthologous genes may be sufficiently related to their closest formal paralog in the same species as to make the evolutionary connection between the two pairs unclear. Nonetheless, it came as a surprise

8 Insect Cytochrome P450

to find a very small number of 1 : 1 pairs of orthologous P450 genes between D. melanogaster and A. gambiae (Ranson et al., 2002a). Zdobnov et al. (2002) indicate a genome wide level of 44–47% orthologous genes, but for the P450 genes only ten orthologs were found, of which five are mitochondrial P450s (Ranson et al., 2002a). When two P450 genes are recognized as orthologs even though the two species have diverged about 250 million years ago, the most likely explanation is that there is a strong evolutionary constraint that has maintained this orthologous relationship. A similar or identical physiological function for the orthologous pair of P450 enzymes may represent such a constraint. Three of the five mitochondrial P450s have indeed a recognized function that is predicted to be identical in Drosophila and A. gambiae: CYP302A1, CYP314A1, and CYP315A1 are hydroxylases of the ecdysteroid biosynthetic pathway (Warren et al., 2002; Petryk et al., 2003; and see Section 4.1.4.1.1). The number of pairs of orthologs between Drosophila and A. gambiae is however higher than the number of P450 enzymes thought to participate in ecdysteroid metabolism (see Chapter 3.3), so that several other insect-specific or dipteran-specific conserved functions are probably carried out by the remaining pairs of orthologs. There are several cases where one gene in one species has two ‘‘orthologs’’ in the other – this is the case when a gene duplication event occurred in just one of the two species after speciation and 250 million years of separate evolutionary history. These cases merit special attention because knowledge of the function of one of the P450s may quickly lead to understanding the function of its alter egos. The comparative analysis of the P450s of the two dipteran species showed that the deficit of true pairs of orthologs (10 versus the predicted 40) was compensated by a most interesting alternative. Orthologous groups of P450 paralogs were seen, i.e., the phylogenetic analysis clearly identified several cases where one ancestral P450 gene underwent several duplication events in each of the two dipteran lineages. The CYP6A cluster on chromosome 5 of the housefly (Cohen and Feyereisen, 1995) and a cluster of CYP6A genes on the right arm of chromosome 2 in Drosophila are probably syntenic, in view of the synteny of these linkage groups (Weller and Foster, 1993), but the clusters have evolved separately for over 100 million years, and orthologous pairs of genes in each cluster can no longer be recognized. Even when syntenic relationships are not maintained globally (Zdobnov et al., 2002), cases of local microsynteny are observed. For example, CYP302A1

and CYP49A1 are very close on chromosome 2L in A. gambiae, but their orthologs in Drosophila are on 3L and 2R, respectively. Nonetheless, microsynteny is maintained around CYP49A1, which is close to the adrenodoxin reductase gene and is located in both species on the negative strand of the first intron of another gene, the G protein 0-a 47A (CG2204) in Drosophila and its ortholog in A. gambiae. 4.1.2.5. Intron/Exon Organization of CYP Genes

The intron/exon organization of P450 genes is a useful tool in the analysis of P450 phylogeny, as shown by the systematic studies in C. elegans (Gotoh, 1998), Arabidopsis thaliana (Paquette et al., 2000), and Drosophila (Tijet et al., 2001) (Figure 3). Intron sequence comparisons, as well as sequence comparisons of 50 flanking sequences have also helped clarify the evolutionary relationships of very closely related CYP6B genes of Papilio species (Li et al., 2002a). Multiple events of intron loss and gain can be deduced from a comparison of the intron/exon organization of orthologous pairs of P450 genes for Drosophila and A. gambiae (Ranson et al., 2002a). Intron phase is nonbiased. CYP introns follow the GT/AG rule, except the first intron in the Drosophila Cyp9c1 gene (Tijet et al., 2001) and the first intron in the Cyp6a8 gene (Maitra et al., 2002). The latter was recognized by comparison with the full length cDNA, it does not conform to usual sequence patterns, and has an AT/TC splice junction. This intron is very short (36 bp) and potentially represents 12 additional in-frame codons (Maitra et al., 2002).

4.1.3. The P450 Enzymatic Complexes 4.1.3.1. Classical Biochemical Approaches

4.1.3.1.1. Subcellular fractions The enzymology of insect P450 can be studied in different types of environments. These are enriched subcellular organelles from insects (microsomes, mitochondria) where multiple P450 interact in situ with their redox partners; membranes from cellular expression systems where a cloned recombinant P450 interacts with native or engineered redox partners; and ultimately a reconstituted system of purified recombinant P450 and its redox partners in a defined system devoid of biological membranes. Transgenic expression of P450 genes is another way to study P450 biochemistry, but also regulation. The classical preparation of microsomal fractions from insect tissue homogenates by differential centrifugation has been described extensively by Hodgson (1985) and Wilkinson (1979). It remains, with minor modifications, the most widely used first step in the biochemical characterization of insect

Figure 3 Neighbor-joining tree (with bootstrap value when different from 1000) and intron position of 83 P450 genes of Drosophila melanogaster. Branch point A indicates the mitochondrial P450 clade. Intron positions are shown schematically in the aligned open reading frames with their phase (| phase 0; [phase 1 and] phase 2). (Reprinted with permission from Tijet, N., Helvig, C., Feyereisen, R., 2001. The cytochrome P450 gene superfamily in Drosophila melanogaster : annotation, intron-exon organization and phylogeny. Gene 262, 189–198. For updated information see [http://P450.antibes.inra.fr].)

10 Insect Cytochrome P450

Figure 4 Sucrose density centrifugation separation of subcellular fractions of housefly larval homogenates showing the distribution of marker enzyme activities between mitochondrial and microsomal fractions. Note some P450 reductase activity in the top (soluble) fractions representing proteolytically cleaved enzyme. (Reprinted with permission from Feyereisen, R., 1983. Polysubstrate monooxygenases (cytochrome P-450) in larvae of susceptible and resistant strains of house flies. Pestic. Biochem. Physiol. 19, 262–269; ß Elsevier.)

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P450 enzymes. Linear or step gradients of sucrose for the centrifugal preparation of microsomes and mitochondria, or CaCl2 precipitation of microsomes have been less favored. In all approaches, the careful use of marker enzymes is critical. A technique for the rapid preparation of microsomal fractions of small tissue samples relying on centrifugation at very high speed on sucrose layers in a vertical rotor has been described (Feyereisen et al., 1985). The well-documented sedimentation of P450associated activities at low g forces in many early insect studies (reviews: Wilkinson and Brattsten, 1972; Wilkinson, 1979) has been considered a peculiar difficulty of insect biochemistry. At the time, the vertebrate toxicology and endocrinology literature had clearly identified P450 metabolism of xenobiotics as microsomal, whereas mitochondrial P450s were chiefly involved in hormone metabolism. There was therefore little incentive in probing the subcellular distribution of insect P450 activity

more carefully. The discovery of insect CYP12 enzymes and their characterization as mitochondrial P450 enzymes capable of metabolizing xenobiotics (Guzov et al., 1998) has shed a new light on the early difficulties in sedimenting insect P450 activities in the ‘‘correct’’ fractions. It is quite probable that at least a part of the P450 activities observed in ‘‘mitochondrial’’ fractions were indeed carried out by CYP12 enzymes. In housefly larvae, 15–20% of the aldrin and heptachlor epoxidase activities were associated with mitochondrial fractions after sucrose density centrifugation (Feyereisen, 1983) (Figure 4). The insect midgut is a particularly rich source of P450 activity (Hodgson, 1985), but the external brush border membrane is a significant source of membrane vesicles (BBMV) upon homogenization of the tissue. Centrifugal methods to separate the BBMV fraction from the microsomes derived from the endoplasmic reticulum have been described (Neal and Reuveni, 1992).

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Figure 6 Type I substrate-induced difference spectrum of recombinant CYP12A1 with increasing concentrations of progesterone. Figure 5 CO-difference spectrum of recombinant CYP12A1. The P450 was reduced with either sodium dithionite (solid line) or with bovine adrenodoxin, adrenodoxin reductase, and NADPH (dashed line).

4.1.3.1.2. Spectral characterization and ligand binding The analysis of P450 levels in subcellular fractions follows the original procedure of Omura and Sato (1964). A difference spectrum between reduced microsomes and reduced microsomes after gentle bubbling of CO readily displays the famous redshifted Soret peak at 450 nm (Figure 5). The concentration of P450 can be calculated from the DOD between 490 and 450 nm and Omura and Sato’s extinction coefficient e ¼ 91 M1 cm1. This measure gives the total concentration of all forms of P450 present in the preparation. Individual P450 proteins may have peaks that are 1 or 2 nm off the 450 nm norm, and when they represent a large portion of the total P450, the total P450 peak may be shifted as a consequence. The degradation of P450 to the inactive P420 form may interfere with the measurement of P450, as already reviewed by Hodgson (1985), and the respiratory chain pigments interfere with the measurement of P450 in mitochondrial fractions. The classical Omura and Sato procedure remains the procedure of choice to measure the amount and purity of P450 proteins produced in heterologous systems (see below). Ligand-induced spectral changes also follow classical procedures detailed in a useful review (Jefcoate, 1978). Type I spectra (peak at 380–390 nm, trough at 415–425 nm) result from ligand in the substrate binding site displacing water as sixth ligand to the heme iron (see Section 4.1.3.4.1, Figure 6). Type I spectra are concentration dependent, giving a spectral dissociation constant (Ks) and this titration is

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correlated with a shift of the iron from low spin to high spin. Not all type I ligands are substrates, and not all substrates are type I ligands, so this useful tool must be used with caution. A type I spectral Ks is not an enzymatic Kd. Type II spectra (peak at 425–435 nm, trough at 390–405 nm) result from the binding of a strong ligand to the heme iron, typically the nitrogen coordination of compounds such as pyrimidines, azoles, or n-octylamine. Type II spectral titration is correlated with a shift from high spin to low spin and is a hallmark of strong inhibitors such as imidazoles (Figure 6). Other, less frequently studied spectral changes induced by ligands or their metabolism will not be discussed here (e.g., type III spectra, Hodgson, 1985; spectra of phenyl–iron complexes, Andersen et al., 1997). 4.1.3.1.3. Assays and substrates Measurement of P450 activity is a special challenge because of the large number of different P450 enzymes, each catalyzing the metabolism of a specific (broad or narrow) range of substrates. There is therefore a very large number of assays for P450 activity. Direct assays of product appearance or substrate disappearance rely on all the tools of analytical chemistry. Indirect assays (e.g., activity of a P450 product in an enzyme or bioassay) can be useful but the strength of the claim for P450 activity depends on the purpose of the assay. The assay of a P450 produced in a heterologous system can be straightforward, but the assay of a P450 in its native microsomal or mitochondrial membrane, where it is mixed with an undetermined number and amount of other P450s, is more problematic. Metabolism of compound M

p0175

12 Insect Cytochrome P450

to product N in microsomes is the sum of the contributions of all P450 enzymes that catalyze the M to N reaction (and sometimes non-P450 enzymes can catalyze the same reaction!). Selective inhibitors (chemicals or antibodies) of one P450 can, by substraction, indicate the relative contribution of that particular P450 to the reaction being measured (e.g., Wheelock and Scott, 1992a; Hatano and Scott, 1993; Korytko et al., 2000b). This indirect inference is only as valid as the inhibitor is selective. Substrates that are selective for one P450 and that are easily assayed have been the object of considerable research in biomedical toxicology. The relative success of this quest (e.g., nifedipine as model probe for CYP3A4) is a result of both the limited number of major P450 expressed in human liver and the heavy investment in their study. The large number of insect P450s times the large number of insect species under study divided by the investment in their research makes a similar quest seem quixotic. By default then, but mostly by inertia of the historical development of insect P450 research, a certain number of assays have taken their place in the literature as some measure of ‘‘global’’ P450 activity. Most authors are now fully aware that the microsomal activity of, e.g., aldrin epoxidation, p-nitroanisole O-demethylation or 7-ethoxycoumarin O-deethylation is only a measure of those P450 enzymes catalyzing these reactions. But this awareness is only as recent as our understanding that there are really many P450 enzymes, and that their individual catalytic competence may be broad or narrow, overlapping with other P450 enzymes or not. Thus, the pioneers who used aldrin epoxidation as an assay did so at a time when P450 research was still strongly influenced by the dichotomy between steroid metabolism by specific P450 enzymes and drug metabolism by two major forms of liver P450. They used an analytical tool then readily available in pesticide toxicology laboratories (GC with electron capture detection for the sensitive detection of organochlorine pesticide residues), but they didn’t use it without a caveat. Quoting the classical Krieger et al. (1971) study: ‘‘to the extent that the rate of epoxidation of aldrin to dieldrin typifies the activity of the enzymes toward a wider range of substrates. . .’’ The current and still widespread use of a ‘‘model’’ substrate to explore P450 activities in insect subcellular fractions can be useful. If the metabolism of a randomly chosen P450 substrate (e.g., aldrin, aminopyrine, 7-methoxyresorufin) is quantitatively different between insect strains, in different tissues, following induction, etc., then this substrate has provided a clue that the qualitative or quantitative complement of P450 enzymes is changing. It is

Figure 7 O-Dealkylation activity of E. coli-expressed recombinant housefly CYP12A1 in a reconstituted system. (Reprinted with permission from Guzov, V.M., Unnithan, G.C., Chernogolov, A.A., Feyereisen, R., 1998. CYP12A1, a mitochondrial cytochrome P450 from the house fly. Arch. Biochem. Biophys. 359, 231–240; ß Elsevier.)

up to the investigator to follow up on this clue. Alkoxycoumarins and alkoxyresorufins are useful substrates for sensitive fluorometric assays and have largely replaced organochlorines as model substrates. They can be used to ‘‘map’’ the catalytic competence of a heterologously expressed P450 (e.g., the preference of CYP12A1 for pentoxycoumarin, Guzov et al., 1998) (Figure 7). Steroids such as testosterone have multiple sites of attack by P450 enzymes and can likewise be used to characterize the activity of subcellular fractions or of heterologously expressed P450s (Amichot et al., 1998; Cuany et al., 1990; M.B. Murataliev, V.M. Guzov, R. Feyereisen, unpublished data). Another approach to the study of P450 activity is to follow the consumption of the other substrates, O2 or NADPH. This approach is certainly valid when the stoichiometry of the reaction (see eqn [1]) is under study, but its use to monitor metabolism by subcellular fractions is fraught with difficulties. Other enzymes consume O2 and NADPH as well, and the background activity can be very high. 4.1.3.1.4. P450 assays in individual insects The design of assays suitable for assessing P450 activities in single insects has accompanied the need to study variations in P450 activities from the individual to the population level. Such assays allow the presentation of frequency histograms of activity levels in field-collected samples or in laboratory-selected populations and are ideally adapted to microtiter plate format. The NADPH-dependent conversion of p-nitroanisole to p-nitrophenol was followed in individual homogenates of H. virescens and

Insect Cytochrome P450 13

Pseudoplusia includens larvae (Kirby et al., 1994; Rose et al., 1995; Thomas et al., 1996). This assay has a relatively low sensitivity, but clearly distinguished individuals from susceptible and insecticide-resistant strains. Cut abdomens of adult Drosophila in buffer containing 7-ethoxycoumarin can be used to measure 7-hydroxycoumarin formation in a 96-well microtiter plate format (de Sousa et al., 1995). This rapid technique allows for instance the monitoring of individual variability over the course of a selection regime (Bride et al., 1997). In another example, 10 000 g supernatants of individual homogenates of Chironomus riparius larvae were assayed for 7-ethoxyresorufin O-deethylation activity (Fisher et al., 2003). The classical aldrin epoxidase assay adapted on single larvae of Spodoptera frugiperda has also been reported, but this assay is not adapted to the 96-well format (Yu, 1991, 1992). A simple assay based on the peroxidase activity of the heme group with tetramethylbenzidine was developed for use in single mosquitoes (Brogdon, 1997). This assay, easily developed on a microtiter plate format, is an indirect assay measuring total heme content of the insect homogenate rather than P450 activity, and therefore needs to be carefully validated. 4.1.3.1.5. Solubilization and purification Solubilization and purification of insect P450 has generally followed the advances pioneered in the purification of vertebrate P450 (Agosin, 1985; Hodgson, 1985) and relatively a few studies since 1985 have pursued this difficult task (Ronis et al., 1988; Wheelock and Scott, 1989). P450 purification from microsomes of mixed tissues (e.g., fly abdomens) can be sufficient to obtain a protein fraction suitable for antibody generation or peptide sequencing (Feyereisen et al., 1989; Wheelock and Scott, 1990; Scott and Lee, 1993b). Sequential chromatography on octylamino agarose, DEAE-cellulose, and hydroxyapatite was used to purify sodium cholate-solubilized P450s from Drosophila (Sundseth et al., 1990). The two protein fractions obtained, P450 A and B, had only a very low 7-ethoxycoumarin O-deethylase activity (0.01 nmol/nmol P450/min), but the proteins were useful in generating monoclonal antibodies that allowed the subsequent cloning of CYP6A2 (Waters et al., 1992). An original approach was the purification of a locust P450 by affinity chromatography with type II and type I ligands (Winter et al., 2001) that led to cloning of CYP6H1 (Winter et al., 1999). In this approach, microsomes from larval Locusta

migratoria Malpighian tubules were first treated with the detergent synperonic NP10 to solubilize P450. The extract was then chromatographed on o-octylamino agarose then hydroxylapatite. The third and less classical step was chromatography on a triazole agarose affinity column. The affinity ligand was a derivative of the fungicide difenoconazole, which has an affinity for ecdysone 20-monoxygenase of the same level as that of the substrate ecdysone (0.5 versus 0.2 mM). This substituted triazole is a typical type II ligand (active site liganding of the heme) and its use on the affinity matrix led to the adsorption of all the P450 loaded on the column (Winter et al., 2001). Elution from the affinity column was done by replacing the immobilized type II ligand with a soluble type I ligand, ecdysone (see Chapter 3.3). A major protein band of 60 kDa was thus obtained in 4% yield, with a P450 specific activity of 13.1 nmol/mg protein. Unfortunately, biochemical evidence that this P450 is in fact an ecdysone 20-monooxygenase was not obtained in this study (Winter et al., 2001), so the nature of CYP6H1 (Winter et al., 1999) remains conjectural. Another variant on the classical purification schemes has been the use of immobilized artificial membrane high performance liquid chromatography (IAM-HPLC) of microsomal proteins (Scharf et al., 1998). This technique allowed the 70-fold purification of a P450 from the German cockroach and the subsequent production of antibodies with this 49 kDa protein as antigen. 4.1.3.2. Heterologous Expression Systems

Biochemical characterization of a P450 protein and its substrate selectivity remains a sine qua non condition of its functional identification. Only a few P450 enzymes are characterized well enough (e.g., steroid metabolizing P450s in vertebrates) that sequence comparison can reasonably predict activity. For most other P450s, the sequence does not provide a clue to the activity, and there are now innumerable papers describing how one or a few mutations can change substrate selectivity (review: in Domanski and Halpert, 2001). Only three mutations are needed to confer to Drosophila CYP6A2 the ability to metabolize DDT (Berge´ et al., 1998; Amichot et al., 2004). In the absence of significant studies on the activity of P450 proteins purified directly from insect tissues, it is the expression of P450 cDNAs in heterologous systems that has become the standard way of characterizing insect P450 proteins. A number of such expression systems have been developed over the last few years (Table 1), and the techniques are essentially similar to those used for the production of P450 proteins from mammalian or plant tissues.

14 Insect Cytochrome P450

t0005

Table 1 Heterologous expression systems for insect P450 Expression system

P450 produced

Substrate metabolized

Reference

Escherichia coli

CYP4C7 CYP6A1

Sutherland et al. (1998) Andersen et al. (1994) Andersen et al. (1997) Sabourault et al. (2001)

CYP15A1

Sesquiterpenoids Aldrin, heptachlor Sesquiterpenoids Diazinon 7-Propoxycoumarin 1-Bromochlordene, chlordene, 1-hydroxychlordene, isodrin Testosterone, progesterone, androstenedione Pisatin Chlorfenapyr DDT, testosterone Benzphetamine, p-chloro-N-methylanilin, methoxyresorufin Sesquiterpenoids Aldrin, heptachlor, diazinon, azinphosmethyl, amitraz, progesterone, testosterone, 7-alkoxycoumarins t t, methyl farnesoate

CYP6A2 CYP6B1

Aldrin, heptachlor, diazinon Furanocoumarins

CYP6B4 CYP6B17 CYP6B21 CYP6B25

Furanocoumarins, ethoxycoumarin Furanocoumarins, ethoxycoumarin Furanocoumarins, ethoxycoumarin Furanocoumarins

Dunkov et al. (1997) Ma et al. (1994), Hung et al. (1997), Wen et al. (2003) Hung et al. (1997), Li et al. (2003) Li et al. (2003) Li et al. (2003) Li et al. (2003)

Yeast

CYP6A2 CYP6D1

AflatoxinB1, 7,12-dimethylbenz[a]anthracene, 3-amino-1-methyl-5H-pyrido[4,3-b]-indole Methoxyresorufin

Smith and Scott (1997)

Transfected S2 cells

CYP302A1 CYP314A1 CYP315A1

2,22-Dideoxyecdysone Ecdysone 2-Deoxyecdysone, 2,22-dideoxyecdysone

Warren et al. (2002) Petryk et al. (2003) Warren et al. (2002)

CYP6A2 CYP6A5 CYP9E1 CYP12A1

Baculovirus

a b

c

d e

Amichot et al. (2004) f

g

Guzov et al. (1998)

Helvig et al. (2004)

Saner et al. (1996)

a

V.M. Guzov and R. Feyereisen, unpublished data. J. Walding, J.F. Andersen, and R. Feyereisen, unpublished data. c M.B. Murataliev, V.M. Guzov, and R. Feyereisen, unpublished data. d V.M. Guzov, H. VanEtten, and R. Feyereisen, unpublished data. e V.M. Guzov, M. Kao, B.C. Black, and R. Feyereisen, unpublished data. f J.L. Stevens, J.F. Andersen, and R. Feyereisen, unpublished data. g J.F. Andersen and R. Feyereisen, unpublished data. b

4.1.3.2.1. Escherichia coli Bacterial production (Escherichia coli) of insect P450s has required several modifications of the sequence. At the 50 end of the cDNA, mutations are introduced to optimize expression (Barnes et al., 1991). The second codon is replaced by Ala (Andersen et al., 1994; Guzov et al., 1998; Sutherland et al., 1998) and silent substitutions are introduced to increase the A/T content (Sutherland et al., 1998). In some cases, introduction of 4–6 His codons just before the stop codon directs the translational production of a C-terminal ‘‘histidine tag’’ (Guzov et al., 1998; Sutherland et al., 1998). P450 production can be enhanced by the addition of d-aminolevulinic acid (a precursor for

heme biosynthesis) to the culture broth (Sutherland et al., 1998). The P450 produced in bacteria is found mostly in a membrane fraction, and sometimes in a fraction of inclusion bodies that are difficult to extract. In some cases (Andersen et al., 1994), a significant amount of P450 is produced as a soluble form. The E. coli membrane fraction carrying the recombinant P450 protein is generally suitable for analysis by difference spectroscopy for either P450 content by the Omura and Sato (1964) procedure, or for ligand binding (type I binding for potential substrates or type II binding for azoles). Although some P450 proteins (e.g., CYP17) expressed in E. coli can utilize an endogenous flavodoxin

Insect Cytochrome P450 15

reductase/flavodoxin system for catalysis, none of the insect P450 proteins tested thus far have been catalytically active in E. coli membrane fractions in the absence of a P450 reductase. Therefore, P450 produced in bacteria needs to be solubilized and purified by classical methods. The proteins produced with a histidine tag, once solubilized, are purified by nickel chelate affinity chromatography. Extensive dialysis is needed in both procedures to remove excess detergent or imidazole used for elution from the nickel affinity column. The P450 obtained is then suitable for reconstitution with redox partners. These partners (microsomal or mitochondrial redox partners, see Section 4.1.3.3) are themselves produced in E. coli and purified (Guzov et al., 1998). Reconstitution of a catalytically active enzyme system is then tedious or artistic, depending on one’s degree of patience. It requires attention to the details of concentrations of phospholipids, detergents, proteins, and their order of addition, mixing and dilution (Sutherland et al., 1998). The advantages of bacterial expression are the low cost of production of large amounts of P450, and the possibility to work with a precisely defined in vitro system with highly purified enzymes and their partners. A thorough characterization of the enzyme can be undertaken. The disadvantage of this formal biochemical approach is that purification and reconstitution are difficult and time-consuming, and is probably not suitable for when the goal is simply a survey of the catalytic competence of the P450, or the comparison of a large number of P450s or P450 mutants. The host organism, E. coli, is a rare organism devoid of P450 genes of its own while other bacteria can carry over 20. 4.1.3.2.2. Baculovirus Expression of P450 in lepidopteran cells by the baculovirus system requires no modification of sequence and is a widely used method for the production of proteins in an eukaryotic system. It has the potential of producing large amounts of P450 proteins for subsequent purification, but studies with insect P450 expressed with this system (Ma et al., 1994; Dunkov et al., 1997; Hung et al., 1997; Chen et al., 2002; Wen et al., 2003) have relied instead on the advantage that the protein is present in a suitable milieu, the endoplasmic reticulum of an insect cell. Thus, cell lysates, briefly centrifuged to pellet cell debris, are used as enzyme source. Difference spectroscopy or immunological methods (Dunkov et al., 1997) can be used to assess the amount of P450 produced. The host cells provide their endogenous P450 reductase to support the activity of the heterologous P450 when the cell lysates are incubated with an NADPH

regenerating system. Although the level of P450 reductase is sufficient to allow the measurement of a number of P450-dependent activities (Dunkov et al., 1997), the stoichiometry of endogenous P450 reductase, and cytochrome b5, to heterologously expressed P450 is probably not optimal. The activities measured do not represent the full potential of the P450 under study. For instance, a thirty-fold increase in CYP6A2 activity was observed when purified housefly P450 reductase and cytochrome b5 were added to lysates of cells expressing Cyp6a2 (Dunkov et al., 1997). An improvement of the baculovirus expression system has therefore been designed, wherein the cells are coinfected with a virus engineered to carry the P450 and a virus engineered to carry a P450 reductase (housefly P450 reductase, Wen et al., 2003). Optimal conditions were sought, and a significant increase (33-fold) in CYP6B1 activity towards the substrate xanthotoxin was achieved with the improved P450 reductase/P450 ratio. In fact, the improved conditions allowed the measurement of angelicin metabolism that was barely detectable in the absence of additional P450 reductase. Thus, in the baculovirus system, insect P450s can be studied in an insect membrane environment, without need for purification. Those are great advantages over the E. coli expression system. However, the interactions with its redox partners are not manipulated as easily (Wen et al., 2003). The total amounts of P450 produced are also smaller, although addition of hemin to the culture medium can increase the amount of P450 produced (Dunkov et al., 1997; Wen et al., 2003). The total amount of P450 produced is less important in the baculovirus system than in the E. coli system as purification is not required for most applications, and as the highest activity of cell lysates is achieved at the correct P450/P450 reductase ratio, not at the maximal P450 production level (Wen et al., 2003). The level of endogenous P450 in the control experiments, i.e., uninfected cells or cells infected with a virus carrying a nonP450 ‘‘control’’ cDNA, are virtually undetectable. 4.1.3.2.3. Transfection in cell lines Heterologous expression in transfected mammalian COS cells was first established in 1986 for bovine CYP17 (Zuber et al., 1986), but it is not until later that an insect P450 was similarly expressed in an insect cell line. Thus, Drosophila Schneider 2 cells have been transfected with Drosophila P450 cDNAs (Warren et al., 2002). Expression under control of the actin 5C promoter produced sufficient P450 for activity measurements. The advantage of the method is its simplicity. When the expression of the P450 is coupled

16 Insect Cytochrome P450

with a very sensitive assay, the method can rapidly provide qualitative data on the catalytic competence of the enzyme. However, the quantitative determination of P450 levels is more difficult to achieve, and the interaction with redox partners cannot be optimized except by coinfection. It is interesting that the CYP302A1 and CYP315A1 expressed by this method are mitochondrial P450s and the S2 cell homogenates were able to provide adequate redox partners. As used so far, it has not allowed a measurement of the amounts of P450 produced, nor have the redox partners been characterized or optimized. Cell transfection does not have the potential of the baculovirus system for large-scale production of P450 proteins. 4.1.3.2.4. Yeast Yeast expression systems have only started to be exploited for the production of insect P450 proteins. Saccharomyces cerevisiae has three P450 genes that are fully characterized, and are expressed at low levels so that inducible expression of an exogenous P450 is not hindered by the endogenous P450. Coproduction of CYP6A2 from Drosophila and of human P450 reductase in yeast (Saner et al., 1996) generated a cell system capable of activating several procarcinogens to active metabolites that induced mitotic gene conversion or cytotoxicity. Housefly CYP6D1 was also produced in yeast but methoxyresorufin demethylation was the only marker activity obtained with microsomes of the transformed yeast (Smith and Scott, 1997). Insect P450 production in yeast has not yet achieved the success seen with plant P450 production in yeast (Schuler and Werck-Reichhart, 2003). P450 cDNAs may need to be engineered to recode the N-terminus of the protein. This has been done successfully with plant P450s to conform with the yeast codon usage (Hehn et al., 2002). The replacement of the yeast P450 reductase gene with an insect P450 reductase gene by homologous recombination (Pompon et al., 1996) should increase the usefulness of this yet underutilized expression system. Indeed, yeast combines the advantages of E. coli inducible production of large amounts of protein with the advantage of the eukaryotic cell system in which P450 enzymes can be studied in a normal membrane environment. 4.1.3.2.5. Transgenic insects The use of transgenic insects to study P450 function (or regulation, see Sections 4.1.4.5.3 and 4.1.5.2.2) has until now been restricted to Drosophila. In the first report, Gandhi et al. were unable to rescue by transgenesis the lethality of two complementation groups in the Cyp4d1 region (Gandhi et al., 1992). Heterologous expression of vertebrate P450s was achieved in

studies aimed at developing Drosophila as a genotoxicity model organism. The rat CYP2B1 gene was expressed under control of the Drosophila LSP1a promoter (Jowett et al., 1991). This promoter ensures high levels of expression in third instar larvae. Transgenic flies expressed functional CYP2B1, as shown by increased CYP2B1-specific metabolism of 7-benzyloxyresorufin and by increased sensitivity to cyclophosphamide, a procarcinogenic drug activated by CYP2B1. In similar experiments, canine CYP1A1 was expressed under the control of the Drosophila heatshock inducible hsp70 promoter. Small amounts of CYP1A1 were produced after heatshock, sufficient to increase the sensitivity of the flies to 7,12-dimethylbenz[a]anthracene, a polycyclic aromatic hydrocarbon that is metabolized by CYP1A1 to a genotoxic metabolite (Komori et al., 1993). Housefly CYP6D1 was produced in Drosophila under control of the heatshock inducible hsp70 promoter, and this led to a significant increase in benzo[a]pyrene hydroxylation (Korytko et al., 2000a), though heat shock decreased total P450 levels in whole body microsomes. Transgenic expression of CYP6G1 has been an important piece of evidence in demonstrating its role in DDT and neonicotinoid resistance (Daborn et al., 2002; Le Goff et al., 2003) as discussed below (Section 4.1.4.5.5). Transformation of other insects (see Chapter 4.13), notably with the piggyBac vector as in Bombyx mori (Tamura et al., 2000) will undoubtedly increase the applications of transgenesis to P450 research. 4.1.3.3. P450 Enzymes and Their Redox Partners

4.1.3.3.1. P450 proteins The sequence identity of distantly related P450 proteins can be as low as that predicted from the random assortment of two sets of 500 or so amino acids. This is because there are very few absolutely conserved amino acids. In insect sequences available to date, these are found in five conserved motifs of the protein (Figure 8), the WxxxR motif, the GxE/D TT/S motif, the ExLR motif, the PxxFxPE/DRF motif and the PFxxGxRxCxG/A motif. Despite this tremendous overall sequence diversity, the increasing number of crystal structures for P450 proteins, mostly soluble forms from bacteria (Poulos et al., 1995), reveals a quite high conservation of the three-dimensional structure. The description of this structure essentially follows the nomenclature of the P450cam protein, the camphor hydroxylase of Pseudomonas putida (Poulos et al., 1985). The first motif WxxxR is located in the C-helix, and the Arg is thought to form a charge pair with the propionate of the heme. This

Insect Cytochrome P450 17

Figure 8 Conserved and variable regions of P450 proteins illustrated over their primary structure (sequence). (Adapted with permission from Werck-Reichhart, D., Feyereisen, R., 2000. Cytochromes P450: a success story. Genome Biol. 1, 3003.1–3003.9; ß GenomeBiology.)

motif is not easily discernible, except in multiple alignments. The second conserved motif GxE/DTT/ S surrounds a conserved threonine in the middle of the long helix I that runs on top of the plane of the heme, over pyrrole ring B. The third conserved motif ExLR is located in helix K. It is thought to stabilize the overall structure through a set of salt bridge interactions (E-R-R) with the fourth conserved motif PxxFxPE/DRF (often PERF, but R is sometimes replaced by H or N) that is located after the K0 helix in the ‘‘meander’’ facing the ExLR motif (Hasemann et al., 1995). The fifth conserved motif PFxxGxRxCxG/A precedes helix L and carries the cysteine (thiolate) ligand to the heme iron on the opposite side of helix I. The cysteine ligand is responsible for the typical 450 nm (hence P450) absorption of the FeII–CO complex of P450 (Mansuy and Renaud, 1995). This heme binding loop is the most conserved portion of the protein, often considered as ‘‘signature’’ for P450 proteins. Deviations from the consensus sequences of these five motifs deserve special attention. For instance, the CYP301A1 of both Drosophila and A. gambiae has a very unusual Tyr instead of Phe in the canonical PFxxGxRxCxG/A motif around the Cys axial ligand to the heme. These deviations may denote an atypical catalytic

function for the P450 enzyme, as seen in P450 enzymes that are not monooxygenases, such as plant allene oxide synthase (CYP74A) or vertebrate thromboxane synthase (CYP5A1) whose I helix lacks the conserved Thr. In the bacterial hydroxylase P450eryF (CYP107A), the Thr is replaced by Ala. A water molecule and a hydroxyl group of the substrate have become functional equivalents of the Thr hydroxyl (Poulos et al., 1995). P450 proteins are also characterized by their Nterminal sequence (Figure 9). Those targeted to the endoplasmic reticulum have a stretch of about 20 hydrophobic amino acids. These precede one or two charged residues that serve as halt-transfer signal and a short motif of prolines and glycines. The latter serves as a ‘‘hinge’’ that slaps the globular domain of the protein onto the surface of the membrane while the N-terminus is anchored through it. The presence of the PGPP hinge is necessary for proper heme incorporation and assembly of functional P450s in the cell (Yamazaki et al., 1993; Chen et al., 1998). A hydrophobic region between helices F and G is thought to penetrate the lipid bilayer, thus increasing the contact of the P450 with the hydrophobic environment from which many substrates can enter the active site (Williams et al., 2000).

18 Insect Cytochrome P450

Figure 9 Scheme of the N-terminal sequence of microsomal and mitochondrial P450 proteins.

The N-terminal sequence of P450 proteins targeted to mitochondria is usually somewhat longer, and shows several charged residues (Figure 9). The mature mitochondrial protein is proteolytically cleaved at a position that has not been formally recognized for insect mitochondrial P450s to date, but is known for several mitochondrial P450s of vertebrate species. Mitochondrial P450 proteins are also characterized by a pair of charged amino acids in the K helix, R391 and K395 in CYP12A1. Homologous amino acids in mammalian P450scc (K377 and K381) are responsible for the high affinity to the ferrodoxin-type (adrenodoxin) electron donor (Wada and Waterman, 1992; Pikuleva et al., 1999). An additional positively charged residue (R454 of CYP12A1) is homologous to R418 of CYP27A1 shown to increase affinity to adrenodoxin even further (Pikuleva et al., 1999). The insect CYP12, 49, 301, 302, 314, and 315 proteins are most closely related to the mammalian mitochondrial P450 (CYP11, CYP24, and CYP27 families), to CYP44A1 from C. elegans, and to the pond snail CYP10. Subcellular localization of CYP12A1 by immunogold histochemistry with antibodies raised against the CYP12A1 protein produced in bacteria established the mitochondrial nature of CYP12A1 (Guzov et al., 1998). There is evidence that some vertebrate microsomal P450s, e.g., CYP1A1 and CYP2E1, are cleaved in vivo of their N-terminal anchor, thus revealing a cryptic mitochondrial targeting sequence, and the shortened protein is enzymatically active in mitochondria (Anandatheerthavarada et al., 1999). Thus, the 1–44 residues of CYP1A1 serve a dual targeting role, with 1–32 targeting the protein to the ER cotranslationally, whereas cleavage and exposure of three basic amino acids in residues 33–44 direct the posttranslational transport to mitochondria (Bhagwat et al., 1999). Whether some

insect microsomal P450 proteins (e.g., CYP314A1, see Section 4.1.4.1.1) behave in this fashion is currently unknown. Interspersed throughout the globular domain of the P450 proteins are six regions with a low degree of sequence similarity, covering about 16% of the total length of the protein (Figure 8). Initially recognized in CYP2 proteins by Gotoh (1992), these are called SRS (substrate recognition sites) and this designation has been generically extended to other P450s. P450 enzymes, whether microsomal or mitochondrial, need to interact with redox partners for their supply of reducing equivalents from NADPH. Figure 10 schematically illustrates the two types of electron transfer complexes thus formed, and the following sections provide a description of the redox partners. 4.1.3.3.2. NADPH cytochrome P450 reductase P450 reductase (EC 1.6.2.4) belongs to a family of flavoproteins utilizing both FAD and FMN as cofactors. These diflavin reductases emerged from the ancestral fusion of a gene coding for a ferredoxin reductase with its NADP(H) and FAD binding domains with a gene coding for a flavodoxin with its FMN domain. This origin of the enzyme was first proposed by Porter and Kasper (1986) based on their analysis of the rat P450 reductase sequence. The fusion is dramatically illustrated by the threedimensional structure of P450 reductase (Wang et al., 1997) where the domains are clearly distinguished (Figure 11). The architecture of this domain fusion has been found in a handful of other enzymes (Murataliev et al., 2004a). In some cases, further fusion with a P450 gene has led to self-sufficient P450 proteins, e.g., the fatty acid hydroxylase of Bacillus megaterium, P450BM3 (Nahri and Fulco, 1986) and of Fusarium oxysporum, P450foxy (Nakayama et al., 1996).

s0155

Insect Cytochrome P450 19

Figure 10 Mitochondrial and microsomal P450 redox partners.

Figure 11 Structure of NADPH cytochrome P450 reductase. Top: evolutionary origin of the FMN (blue), FAD and NADP(H) (green) binding domains of the protein (yellow: membrane anchor; gray: connecting domain). Bottom: three-dimensional structure of the enzyme with the domains identified by color. The bound substrate NADPH (red) and cofactors FAD and FMN (yellow) are indicated. (Reprinted with permission from Murataliev, M.B., Feyereisen, R., Walker, F.A., 2004a. Electron transfer by diflavin reductases. Biochem. Biophys. Acta 1698, 1–26; ß Elsevier.)

The insect P450 reductases sequenced to date are clearly orthologous to the mammalian P450 reductases, with an overall amino acid sequence identity of 54% for the housefly P450 reductase, first cloned and sequenced in 1993 (Koener et al., 1993). The housefly P450 reductase gene codes for a protein of 671 amino acids, and was mapped to chromosome III. The P450 reductases of other

insects are very similar to the housefly enzyme – 82% identity for the D. melanogaster enzyme (Hovemann et al., 1997), 57% identity for the Bombyx mori enzyme (Horike et al., 2000), and 75% identity for the A. gambiae P450 reductase (Nikou et al., 2003). The insect, mammalian, and yeast enzymes are functionally interchangeable in reconstituted systems of the purified proteins or in heterologous expression systems. However, no detailed study has documented how well a mammalian or yeast P450 reductase can support the activity of an insect P450 when compared to the cognate insect P450 reductase. Early attempts to purify and characterize the enzyme from microsomes of housefly abdomens were hampered by the facile proteolytic cleavage of the N-terminal portion of the protein. This hydrophobic peptide anchors the reductase in the membrane, and its removal abolishes the ability of the remainder of the protein (‘‘soluble’’ or ‘‘tryptic’’ reductase) to reduce P450s. The proteolytically cleaved reductase nonetheless retains the ability to reduce artificial electron acceptors such as cytochrome c, DCPIP or ferricyanide (Hodgson, 1985). Heterologous expression of the cloned P450 reductase has been achieved in E. coli (Andersen et al., 1994) and in the baculovirus expression system (Wen et al., 2003) and a purification scheme (Murataliev et al., 1999) has produced quantities of enzyme sufficient for a detailed catalytic characterization of the enzyme’s functioning and of its reconstitution with redox partners (Figure 12). P450 reductase is an obligatory partner of microsomal P450 enzymes. Antisera to Spodoptera eridania or housefly P450 reductase inhibit all P450-dependent activities tested (Crankshaw et al.,

20 Insect Cytochrome P450

Figure 12 Reduction of housefly CYP6A1 (a) and cytochrome b5 (b) by NADPH cytochrome P450 reductase. The recombinant proteins expressed in E. coli were reconstituted in vitro. On the left, the kinetics of reduction measured by stopped-flow spectrophotometry are shown with the calculated first-order rate constants. On the right, the end point difference spectra of CYP6A1 and cytochrome b5 after reduction by P450 reductase (solid line) and sodium dithionite (dotted line). (Adapted from Guzov, V.M., Houston, H.L., Murataliev, M.B., Walker, F.A., Feyereisen, R., 1996. Molecular cloning, overexpression in Escherichia coli, structural and functional characterization of house fly cytochrome b5. J. Biol. Chem. 271, 26637–26645.)

1981; Feyereisen and Vincent, 1984). Immunoinhibition with P450 reductase antibodies serves as a strong indication of microsomal P450 involvement in NADPH-dependent activities, such as ecdysone 20-hydroxylation in the cockroach and in B. mori eggs (Halliday et al., 1986; Horike and Sonobe, 1999; Horike et al., 2000) and (Z)-9-tricosene biosynthesis in the housefly (Reed et al., 1994). P450 reductase immunoinhibition could serve as a tool to distinguish P450 dependent activities from flavin monooxygenase (FMO) activities in microsomes from insect sources. P450 reductase also transfers electrons to cytochrome b5 (see below) and to other microsomal enzymes such as heme oxygenase.

p0300

4.1.3.3.3. Cytochrome b5 In contrast to P450 reductase, the role of cytochrome b5 as partner in P450 dependent reactions is considerably more complex (see Section 4.1.3.3.3). The housefly cytochrome b5 is a 134 amino acid protein (Guzov et al., 1996) with 48% sequence identity with the orthologous rat cytochrome b5. Its N-terminal domain of about 100 residues is the heme-binding domain that is about 60% identical to that of the vertebrate

cytochrome b5. Its C-terminal portion is a hydrophobic membrane anchor. A probable fatty acid desaturase-cyt b5 fusion protein has been misidentified as the Drosophila cytochrome b5 (Kula et al., 1995; Scott, 1999; Kula and Rozek, 2000), but the correct ortholog is 76% identical to the housefly cytochrome b5. The H. armigera cytochrome b5 (Ranasinghe and Hobbs, 1999a) is 127 amino acids in length and 51% identical to the housefly cytochrome b5, and the A. gambiae cytochrome b5 is 54% identical (Nikou et al., 2003). The known insect cytochrome b5 sequences differ at their C-terminal from both the vertebrate microsomal and outer mitochonrial membrane cytochrome b5 sequences, so that inferences about the subcellular targeting of the insect protein (Wang et al., 2003) would seem premature. The housefly cytochrome b5 protein was produced in E. coli, purified and fully characterized (Guzov et al., 1996). Absorption spectroscopy and EPR revealed properties very similar to cytochromes b5 from vertebrates. NMR spectra indicated that the orientation of the heme in the protein relative to its a, g meso axis is about 1 : 1. This means that the

Insect Cytochrome P450 21

protein is present in two forms of approximately equal abundance, that result from two modes of insertion of the noncovalently bound heme in the protein between the two coordinating histidines (face up and face down). Expression of the hemebinding domain in E. coli revealed that the heme is kinetically trapped in a 1.2 : 1 ratio of the two isomers, and that this orientation results from the selective binding of heme by the apoprotein (Wang et al., 2003). A redox potential of 26 mV was measured by cyclic voltammetry on a treated gold electrode in the presence of hexamminechromium(III) chloride, and was verified by classical electrochemical titration. Stopped flow spectrophotometry showed that the cytochrome b5 is reduced by housefly P450 reductase at a high rate (5.5 s1) (Guzov et al., 1996) (Figure 12). Cytochrome b5 can also be reduced by its own reductase, an NADH-dependent FAD flavoprotein, and can therefore provide either NADHor NADPH-derived electrons to P450 enzymes. NADH-cytochrome b5 reductase (EC 1.6.2.2) has been studied in Ceratitis capitata and M. domestica (Megias et al., 1984; Zhang and Scott, 1996a). The N-terminus sequence of the purified housefly enzyme aligns to an internal sequence of the Drosophila enzyme (CG5946) indicating that it represents a proteolytically processed form. NADHcytochrome b5 reductase and cytochrome b5 are also known to provide electrons to other acceptors, such as fatty acid desaturases and elongases. 4.1.3.3.4. Redox partners of mitochondrial P450 The redox partners of mitochondrial P450s are adrenodoxin reductase, an NADPH-dependent FAD flavoprotein and adrenodoxin, a [2Fe-2S] ferredoxin-type iron sulfur protein. These are named for the two redox partners of mammalian adrenal mitochondrial P450s, and this designation has been liberally bestowed on proteins from animals that don’t have adrenals. Insect adrenodoxin reductase and adrenodoxin have not been functionally characterized, but their bovine orthologs are capable of supporting the activity of an insect mitochondrial P450, housefly CYP12A1 (Guzov et al., 1998). The reduction of CYP12A1 is rapid and efficient with bovine adrenodoxin reductase/adrenodoxin while under the same conditions housefly microsomal P450 reductase is only marginally effective. A fragment of a Drosophila adrenodoxin-like open reading frame is in GenBank on a stretch of DNA that also encodes a heatshock gene at 67B on the right arm of chromosome 3 (Pauli and Tonka, 1987). This stretch of 95 amino acids is similar (about 46%) to vertebrate adrenodoxin, and was initially identified

as the Drosophila adrenodoxin ortholog (GenBank X06542). But the correct adrenodoxin ortholog was revealed by the complete genome sequence at 64B1 as a 152 amino acid protein, 45% identical to the bovine protein. EPR spectroscopic evidence for the presence of an adrenodoxin-like protein in fat body mitochondria of Spodoptera littoralis has been presented (Shergill et al., 1995). The Drosophila and A. gambiae adrenodoxin reductase and adrenodoxin genes have been annotated. The Drosophila P-element induced mutant dare1 for defective in the avoidance of repellents was found to encode Drosophila adrenodoxin reductase (Freeman et al., 1999). Strong dare mutants undergo developmental arrest, and this phenotype is largely rescued by feeding 20-hydroxyecdysone. Decreasing by half the wild-type expression of dare blocks the olfactory response. The gene is expressed at low levels in all tissues of the adult fly, including the brain and the antennae. Highest expression is found in the prothoracic gland portion of the ring gland of third instar larvae, as well as in the nurse cells of adult ovaries. These tissues are known to require mitochondrial P450s for ecdysteroid production. The 55 kDa protein encoded by dare is 42% identical to the human enzyme. 4.1.3.4. Catalytic Mechanisms

4.1.3.4.1. P450 reactions Little work on insect P450 has focused on the catalytic cycle, and the mechanism derived from our understanding of the bacterial and mammalian P450 enzymes (Ortiz de Montellano, 1995b; Schlichting et al., 2000) will be briefly summarized (Figure 13). The oxidized P450 is a mixture of two forms: a low spin (FeIII) form with water as the sixth coordinated ligand on the opposite side of the Cys thiolate ligand, and a high spin (FeIII) pentacoordinated form. Substrate binding displaces water from the sixth liganding position, leading to a shift to high spin. This shift can be observed (type I spectrum) and is accompanied by a decrease in the redox potential of P450. The P450– substrate complex receives a first electron from a redox partner (P450 reductase or adrenodoxin), and ferrous P450 (FeII) then binds O2. At this step CO can compete with O2 for binding to P450, its binding leads to a stable complex, with absorption maximum at 450 nm (Figure 5), that is catalytically inactive. CO can be displaced by light irradiation at 450 nm. The P450–O2–substrate complex in the form of a ferric peroxide complex then accepts a second electron (from P450 reductase or in some cases cytochrome b5, or from adrenodoxin). Different types of activated oxygen forms of the same

22 Insect Cytochrome P450

Figure 13 Catalytic cycle of P450 enzymes in monooxygenation reactions. Three possible forms of the activated oxygen species are shown. See text for details. Other reactions (reduction, isomerization, dehydration) can be catalyzed by oxygen-free forms of the enzyme. (Adapted with permission from Werck-Reichhart, D., Feyereisen, R., 2000. Cytochromes P450: a success story. Genome Biol. 1, 3003.1–3003.9; ß GenomeBiology.)

P450 enzyme can then be formed, depending on the protonation state of the complex and on the homolytic or heterolytic cleavage of the reduced dioxygen. Although the formal reaction is the insertion of an atom of oxygen into the substrate, the other atom being reduced to water (hence the term ‘‘mixedfunction oxidase’’), the nature of the oxidizing species can vary. A P450 (FeIII O O)2 peroxo-iron w w form, a P450 (FeIII O OH) hydroperoxo form, V w w and a P450 (Fe ¼O) or P450 iron-oxo form are the preferred descriptions of the activated oxygen forms (Ortiz de Montellano, 1995b; Schlichting et al., 2000; Newcomb et al., 2003). The type of reaction catalyzed then depends on the substrate and substrate binding site and varies from hydroxylation to epoxidation, O-, N-, and S-dealkyation, N- and S-oxidations, or at least 60 different chemical reactions. The types of reactions currently known to be catalyzed by insect P450 enzymes are listed in Table 2. The P450(FeII)–substrate complex can function as a reductase, and the P450–O2–substrate complex can also function as an oxidase, releasing superoxide, hydrogen peroxide, or water. Under experimental conditions, NADPH and

molecular oxygen can be substituted by organic hydroperoxides, sodium periodate, etc., in what is called the peroxide shunt (Ortiz de Montellano, 1995a). The obligatory role of P450 reductase in catalysis of the microsomal P450 has been proven in reconstitution experiments, but the role of phospholipids is less clear and has not been specifically studied. The role of cytochrome b5 is discussed below. Activity of the mitochondrial CYP12A1 also showed absolute dependence on reconstitution in the presence of (bovine) mitochondrial redox partners (Guzov et al., 1998). The stoichiometry (eqn [1]) RH þ O2 þ NADPH þ Hþ ! ROH þ H2 O þ NADPþ that commonly describes the monooxygenase (sensu Hayaishi) or mixed-function oxidation (sensu Mason) reaction of P450 has not been confirmed experimentally for any insect P450. A more complex stoichiometry would take into account the ‘‘leakage’’ of activated oxygen species as superoxide, hydrogen

p0340

Insect Cytochrome P450 23

Table 2 Enzymatic reactions catalyzed by insect P450 enzymes Reaction catalyzed

Oxidase activity O2 to H2O, H2O2, O. 2 Monooxygenations Aliphatic hydroxylation C–H hydroxylation O-dealkylation Dehalogenation Epoxidation Aromatic hydroxylation Heteroatom oxidation and dealkylation Phosphorothioate ester oxidation N-dealkylation N-oxidation S-oxidation Aldehyde oxidation Complex and atypical reactions Carbon–carbon cleavage Decarbonylation with C–C cleavage Aromatization Dehydrogenation Dehydration Aldoxime dehydration Reduction Endoperoxide isomerization

P450

CYP6A1 (and probably most P450)

CYP4C7, CYP6A1, CYP6A2, CYP6A8, CYP12A1, CYP302A1, CYP312A1, CYP314A1, CYP315A1 CYP6A1, CYP6D1a, CYP12A1, CYP6A5, CYP6B4, CYP6B17, CYP6B21 CYP6A2 (DDT to DDA, DDD) CYP6A1, CYP6A2, CYP12A1, CYP15A1, CYP9E1 CYP6D1a CYP6A1, CYP6A2, CYP12A1, CYP6D1a CYP12A1, CYP6A5 þ (nicotine) þ (phorate) þ (C-26 hydroxyecdysteroids) þ? (sterols, ecdysteroid) þ (P450hyd) þ (defensive steroids) þ (cholesterol) þ (R-CN biosynthesis)  

a Inference from immunoinhibition experiments. þ, indicates metabolism by microsomal P450, but specific enzyme not identified (substrate indicated), , indicates no evidence to date.

peroxide, and water at the expense of NADPH during catalysis. In a ‘‘well coupled’’ reaction as in (eqn [1]) these by-products would not be formed. It is assumed, but not generally proven, that a specialized P450 metabolizing its favorite substrate would follow stoichiometry (eqn [1]). An approximate balance for CYP6A1 epoxidation of heptachlor (Hept) gives the following results: (M.B. Murataliev, V.M. Guzov, R. Feyereisen, unpublished data). 1 Hept þ 13 O2 þ 11:2 NADPH ! 1 Hept epoxide þ1:5 H2 O þ 11:2 NADPþ þ 9:8 H2 O2

½2

and for testosterone (Tst) hydroxylation under the same experimental conditions: 1 Tst þ 6 O2 þ 4:7 NADPH ! 1 Tst-OH þ 0:5 H2 O þ 4:7 NADPþ þ 4:2 H2 O2

presence or absence of cytochrome b5 (see below). Note that those stoichiometries are not balanced, reflecting experimental error in the measurements and that the addition of superoxide dismutase did not change the amount of H2O2, indicating either no superoxide production, or lack of success in measuring it. The uncoupling of monooxygenation is highly likely to be a common feature of insect P450 enzymes that metabolize xenobiotics of synthetic or plant origin. The generation of reactive oxygen species is a corrolary of active P450 metabolism.

½3

In eqns [2] and [3], the parameters that were measured are underlined. These stoichiometries show that a P450 such as CYP6A1 can be simultaneously an oxidase and a monooxygenase. These coupling stoichiometries (M.B. Murataliev, V.M. Guzov, R. Feyereisen, inpublished data) are dependent on the ratio of P450 and P450 reductase, as well as on the

4.1.3.4.2. P450 reductase 4.1.3.4.2.1. P450 interaction with P450 reductase As seen above, the proper functioning of P450 enzymes depends on an efficient electron supply. In insect microsomes, the ratio of P450 enzymes to P450 reductase is about 6–18 to 1 (Feyereisen et al., 1990). In this ratio, all P450 enzymes are summed, so that the actual ratio of one specific P450 enzyme to P450 reductase is probably smaller. The rate of the overall microsomal P450 reaction (two transfers of one electron) is relatively slow so that dissociation of the P450–P450 reductase complex is possible between the first and the second electron transfer. Indeed, cytochrome b5 can replace P450 reductase

p0345

24 Insect Cytochrome P450

for the supply of the second electron in some cases (see below). The effect of varying the P450/P450 reductase ratio on catalytic rates was measured in a reconstituted system for heptachlor epoxidation by CYP6A1. The rate of epoxidation was determined by the concentration of the binary complex of P450 and P450 reductase, with the same high rate being observed in the presence of an excess of either protein (Figure 14). The half-saturating concentration of either protein was about 0.1 mM in the presence of cytochrome b5 (M.B. Murataliev, V.M. Guzov, R. Feyereisen, unpublished data). This is in good agreement with the Km of 0.14 and 0.5 mM for P450 reductase in the presence and absence of cytochrome b5 measured previously (Guzov et al., 1996). Coinfection of Sf9 cells with baculoviruses carrying CYP6B1 and P450 reductase has revealed that highest catalytic activity was achieved at an equivalent, moderate, multiplicities of infection for the two viruses (Wen et al., 2003). Higher enzymatic activities of cell lysates towards furanocoumarins was not achieved when either protein was produced in excess. This result can be explained in part by documented limitations of the cell’s ability to host, fold, and provide cofactors for both P450 and reductase (Wen et al., 2003), but it also supports the idea that highest activity is achieved for the

f0070

Figure 14 Heptachlor epoxidation by E. coli-expressed recombinant housefly CYP6A1 and NADPH cytochrome P450 reductase. A reconstituted system containing variable concentration of CYP6A1 (n) or P450 reductase (,) and a fixed concentration (0.05 mM) of the reciprocal partner, and a cytochrome b5 concentration of 1.0 mM was incubated in the presence of NADPH. (M.B. Murataliev, V.M. Guzov, R. Feyereisen, unpublished data)

highest concentration of the binary complex of the two partners. 4.1.3.4.2.2. P450 reductase functioning P450 reductase accepts two electrons from NADPH; more precisely, it accepts a hydride ion (one hydrogen plus one electron), and donates two electrons, one at a time, to P450 enzymes. P450 reductase is therefore an enzyme with two substrates: NADPH and the electron acceptor (P450 or artificial acceptor such as cyt c), and two products: NADPþ and the reduced electron acceptor. With two bound flavins and a pathway of electron transfer NADPH > FAD > FMN > P450 (or cyt c), its reduction state during catalysis can theoretically vary between the fully oxidized state (0 el.) and the fully reduced state (4 el.). Studies with the purified recombinant housefly P450 reductase (Murataliev et al., 1999; Murataliev and Feyereisen, 1999; Murataliev and Feyereisen, 2000; review: Murataliev et al., 2004a) have shed light on two questions posed by this electron transfer function: what is the kinetic mechanism of this two-substrate enzyme (Ping-Pong or sequential Bi-Bi) and what are the respective reduction states of the two flavins during catalysis? In the ping-pong mechanism, the first product of the reaction must be released before the second substrate binds to the enzyme, and no ternary complex is formed. In sequential Bi-Bi mechanisms both substrates bind to the enzyme to form a ternary complex. Although several kinetic mechanisms have been proposed (Hodgson, 1985), a careful study of the recombinant housefly P450 reductase clearly established a sequential random Bi-Bi mechanism (Murataliev et al., 1999). The great sensitivity of the enzyme to ionic strength hampers the comparison of different studies (Murataliev et al., 2004a). The formation of a ternary complex of NADPH, P450 reductase, and the electron acceptor suggested a role for reduced nucleotide binding in the catalysis of fast electron transfer. The rate of cytochrome c reduction was shown to equal the rate of hydride ion transfer from the nucleotide donor to FAD (Murataliev et al., 1999). A faster electron transfer rate was observed with NADPH as compared to NADH (Murataliev et al., 1999) and the 20 -phosphate was shown to contribute to more than half of the free energy of binding (Murataliev and Feyereisen, 2000). The affinity of the oxidized P450 reductase was ten times higher for NADPH than for NADPþ (Murataliev et al., 1999), and a conformational change induced by NADPH binding and important for fast catalysis was suggested by these studies. The state of reduction of the flavins of P450 reductase during catalysis was deduced from kinetic

Insect Cytochrome P450 25

Figure 15 Reduction state of NADPH cytochrome P450 reductase during catalysis, the ‘‘0-2-1-0’’ cycle. The enzyme cycles between a fully oxidized state and a 2-electron reduced state. The electron acceptor (A and A0 ¼ P450 or cytochrome c) receives one electron at a time from an FMN semiquinone form (FMN ) of the enzyme. The release of NADPþ is shown here to occur at the last step but may occur earlier. Implicit additional steps are not shown, for clarity. See text for details and Murataliev et al. (2004a) for review.

experiments, rates of NADPH oxidation and EPR measurements of flavin semiquinone (free radical) levels. These revealed the existence of a catalytically competent FMN semiquinone, different from the ‘‘blue’’ neutral FMN semiquinone, known as the air-stable semiquinone that is not a catalytically relevant form of the enzyme (Murataliev and Feyereisen, 1999). Furthermore, the detailed studies of housefly P450 reductase led to a proposed catalytic cycle where the reduction state of the enzyme does not exceed 2 el., and where an FMN semiquinone, and not an FMN hydroquinone, serves as electron donor to the acceptor P450 or cytochrome c. This ‘‘0-2-1-0 cycle’’ (Figure 15) likely represents the general mechanism of P450 reductases, with strong evidence that it operates in P450BM3 and in the human P450 reductase as in the fly P450 reductase (Murataliev et al., 2004a). 4.1.3.4.3. Role of cytochrome b5 Depending on the P450 enzyme and on the reaction catalyzed, cytochrome b5 may be either inhibitory, without effect, or its presence may be obligatory. Cytochrome b5 can have a quantitative effect on overall reaction rates, and/or a qualitative role on the type of reaction catalyzed and the ratio of the reaction products. The role of cytochrome b5 may or may not depend on its redox (electron transfer) properties. It can also influence the overall stoichiometry of

the P450 reaction, in particular the ‘‘coupling rate,’’ i.e., the utilization and fate of electrons from NADPH relative to monooxygenation. Cytochrome b5 should therefore be regarded as an important modulator of microsomal P450 systems. General reviews of the role of cytochrome b5 in P450 reactions are available (Porter, 2002; Schenkman and Jansson, 2003) and known examples of this modulator role in insect systems follow. The relative contribution of NADH in P450 reactions, but more importantly the NADH synergism of NADPH-dependent reactions that is occasionally observed (e.g., Ronis et al., 1988; Feng et al., 1992), is probably attributable to cytochrome b5 as redox partner. Indeed, the Km of the P450 reductase for NADH is a thousand-fold higher than for NADPH, and the Vmax tenfold lower (Murataliev et al., 1999), so that the contribution of NADH under normal conditions is probably channeled by NADH-cytochrome b5 reductase and cytochrome b5. An anticytochrome b5 antiserum severely inhibited (up to 90%) methoxycoumarin and ethoxycoumarin O-dealkylation and benzo[a]pyrene hydroxylation, but not methoxyresorufin and ethoxyresorufin O-dealkylation when assayed in microsomes of the housefly LPR strain (Zhang and Scott, 1994). This antiserum also inhibits cypermethrin 40 -hydroxylation by these CYP6D1-enriched microsomes (Zhang and Scott, 1996b). Housefly cytochrome b5 stimulates heptachlor epoxidation and steroid hydroxylation when reconstituted with cytochrome P450 reductase, housefly CYP6A1, and phospholipids (Guzov et al., 1996; Murataliev et al., 2004a). Stimulation of cyclodiene epoxidation and diazinon metabolism were also observed with Drosophila CYP6A2 expressed with the baculovirus system (Dunkov et al., 1997). Cytochrome b5 is efficiently reduced by P450 reductase (Figure 12), but it does not increase the rate of P450 reduction by P450 reductase. Because of its small redox potential (see above), cytochrome b5 is unlikely to play an important role in delivering the first electron to P450 catalysis, and its stimulatory role probably involves an increased rate of transfer of the second electron. Cytochrome b5 decreases the apparent Km for P450 reductase and increases the Vmax for epoxidation at constant CYP6A1 concentrations (Guzov et al., 1996). The results suggest a role for cytochrome b5 in the P450 reductase–P450 interactions. Whereas heptachlor epoxidation by CYP6A1 was increased two- to threefold by the addition of cytochrome b5, the hydroxylation of testosterone, androstenedione, and progesterone was stimulated

p0385

26 Insect Cytochrome P450

p0390

seven- to tenfold. The addition of cytochrome b5 increased the ratio of 2b-hydroxylation over 15bhydroxylation of testosterone. This suggests that cytochrome b5 can have an effect on CYP6A1 conformation, probably altering the interaction of the binding site with either the C-17 hydroxyl group (decreased) or the C-3 carbonyl (increased). Interestingly, the effect of cytochrome b5 on hydroxylation regioselectivity was also obtained with apo-b5 (cytochrome b5 depleted of heme and therefore redox incompetent), whereas the effect on turnover number was only much smaller with apo-b5. The effect of apo-b5 is not due to heme transfer from P450 to apo-b5 and, in fact, both apo-b5 and (holo) cytochrome b5 were shown to stabilize the ferrous– CO complex of CYP6A1, decreasing the rate of its conversion to P420 (M.B. Murataliev, V.M. Guzov, R. Feyereisen, unpublished data). Cytochrome b5 increases the coupling stoichiometry of CYP6A1 catalysis. In the heptachlor epoxidation assay, coupling (NADPH or O2 used/ heptachlor epoxide formed) increased from 900 ESTs from the CA of vitellogenic females yielded three ESTs matching a P450 sequence by BLAST analysis. A full-length cDNA of this P450 was expressed in E. coli (Helvig et al., 2004). This P450 termed CYP15A1, has a high affinity for methyl farnesoate, showing a type I spectrum with a Ks of 6 mM. The enzyme, when reconstituted with fly P450 reductase, catalyzed the NADPH-dependent epoxidation of 2E, 6E-methyl farnesoate to JH III. The epoxidation is highly stereoselective (98 : 2) to the natural 10R enantiomer over its diastereomer. The enzyme also has a high substrate specificity, epoxidizing the 2E,6E isomer preferentially over the 2Z,6E isomer, and accepting no other substrate tested including farnesoic acid. CYP15A1 is expressed selectively in the CA, and the rank order of inhibition of CYP15A1 activity by substituted imidazoles is

identical to that of JH biosynthesis by isolated CA (Helvig et al., 2004). 4.1.4.1.2.2. CYP4C7 – v-hydroxylase Expression of the CYP15A1 gene is high when JH synthetic levels are high, but another P450 gene, CYP4C7, also selectively expressed in the CA of D. punctata (Sutherland et al., 1998), has an expression pattern that mirrors the pattern of JH synthesis. The recombinant CYP4C7 enzyme produced in E. coli metabolized a variety of sesquiterpenoids but not mono- or diterpenes. In addition to metabolizing JH precursors, farnesol, farnesal, farnesoic acid, and methyl farnesoate, it also metabolized JH III to a major metabolite identified as (10E)-12hydroxy-JH III (Sutherland et al., 1998). Although this o-hydroxylated JH III has not been identified as a product of the CA, L. migratoria CA are known to produce several hydroxylated JHs in the radiochemical assay in vitro (Darrouzet et al., 1997; Mauchamp et al., 1999). These hydroxy-JHs, 80 -OH-, 120 -OH (Darrouzet et al., 1997), and 40 -OH-JH III (Mauchamp et al., 1999) may be major products of the CA after JH III itself and their role is unknown. Their presence suggests that locust CA have a P450 homologous to cockroach CYP4C7 that has a lower regioselectivity. Indeed, the hydroxy-JHs can be synthesized by locust CA from JH III, and this hydroxylation is inhibited by CO and piperonyl butoxide (Couillaud et al., 1996). The tight physiological regulation of CYP4C7 expression in adult female D. punctata (Sutherland et al., 2000) indicates that the terpenoid o-hydroxylase has an important function to play at the end of vitellogenesis and at the time of impending chorionation and ovulation. It was hypothesized (Sutherland et al., 1998, 2000) that this hydroxylation was a first step in the inactivation of the very large amounts of JH and JH precursors present in the CA of this species after the peak of JH synthesis. The study of JH metabolism has long been dominated by esterases and epoxide hydrolases (see Chapter 3.7), but early work on insecticide-resistant strains of the housefly revealed oxidative metabolism as well (review: Hammock, 1985). Evidence that housefly CYP6A1 efficiently metabolizes sesquiterpenoids, including JH to its 6,7-epoxide, confirms these early studies (Andersen et al., 1997). Hydroxylation and epoxidation of JHs are thus confirmed P450-mediated metabolic pathways for these hormones. A comparison of four P450 enzymes that metabolize methyl farnesoate (Table 3) shows their

Insect Cytochrome P450 33

t0015

Table 3 Specificity of insect P450 enzymes towards methyl farnesoate (MF) P450

Substrate

Reaction

Product formed

Reference

CYP15A1 CYP4C7 CYP9E1

2E, 6E-MF Sesquiterpenoids 2Z > 2E-MF (1.5 : 1) MF or JH all 4 isomers

Epoxidation o-Hydroxylation Epoxidation

10R-epoxide 12E-OH 10-Epoxide 10R : 10S (1 : 1) 6 or 10-epoxide 10S : 10R (3 : 1)

Helvig et al. (2004) Sutherland et al. (1998)

CYP6A1

Epoxidation

a

Andersen et al. (1997)

a J.F. Andersen, G.C. Unnithan, J.F. Koener, and R. Feyereisen, unpublished data. Data from Helvig, C., Koener, J.F., Unnithan, G.C., Feyereisen, R., 2004. CYP15A1, the cytochrome P450 that catalyzes epoxidation of methyl farnesoate to juvenile hormone III in cockroach corpora allata. Proc. Natl Acad. Sci. USA 101, 4024–4029; ß National Academy of Sciences, USA.

catalytic versatility. One is extremely substrate specific (CYP15A1), another is not (CYP6A1). Three of them are epoxidases, one is a hydroxylase on this substrate (CYP6A1 also has activity as a hydroxylase – but not on this substrate). One is a stereoselective epoxidase (CYP15A1), another lacks product enantioselectivity (CYP9E1). 4.1.4.1.3. Biosynthesis of long-chain hydrocarbons The cuticle of insects can be characterized by its hydrocarbon composition, and this can serve as a subtle tool in chemical taxonomy (Lockey, 1991). In some insects, cuticular or exocrine alkanes and alkenes can serve as allomones, or even pheromones (see Chapter 3.14). To cite but one example, the Dufour gland secretions of the leaf-cutting ant Atta laevigata are deposited on foraging trails. This trail pheromone comprises n-heptadecane, (Z)-9-nonadecene, 8,11-nonadecadiene, and (Z)-9-tricosene (Salzemann et al., 1992). Despite their apparently simple structure, the biosynthesis of these hydrocarbons is complex and the enzymes involved are still under intense study. Schematically (Figure 19), long chain (18–28 carbons) fatty acid CoA esters are first reduced to their aldehyde by an acyl-CoA reductase, and they are subsequently shortened to form Cn–1 hydrocarbons (Reed et al., 1994). The conversion of (Z)-tetracosenoyl-CoA (from a C24 : 1 fatty acid) to (Z)-9-tricosene (a C23 : 1 alkene) has been characterized in housefly epidermal microsomes. The role of the aldehyde tetracosenal as an intermediate is evidenced by trapping experiments with hydroxylamine. The next step is NADPH- and O2-dependent, and is truly an oxidative decarbonylation reaction, which releases the terminal carbon as CO2 and not as CO (Reed et al., 1994). Inhibition by carbon monoxide and antiP450 reductase antibodies are strongly indicative of a P450 reaction, and this most peculiar enzyme was called P450hyd (Reed et al., 1994). The reaction does not involve a terminal desaturation, because both C-2 protons of the aldehyde are retained in

the hydrocarbon product, and the aldehydic proton is transfered to the product (Reed et al., 1995). NADPH and molecular oxygen can be replaced by ‘‘peroxide shunt’’ donors, suggesting that this reaction involves a perferryl iron-oxene species of activated oxygen (Ortiz de Montellano, 1995a), and that it proceeds through an alkyl radical intermediate (Reed et al., 1995). This type of oxidative decarbonylation reaction may be widespread in insects because a mechanistically identical conversion of octadecanal to heptadecane has been documented in flies, cockroaches, crickets, and termites (Mpuru et al., 1996). 4.1.4.1.4. Pheromone metabolism In female houseflies, the (Z)-9-tricosene produced by P450hyd is a major component of the sex pheromone, being responsible for inducing the courtship ritual and the males’ striking activity (see Chapter 3.14). Additional components of the pheromone are the sex recognition factors (Z)-9,10-epoxytricosane and (Z)-14-tricosen-10-one. These compounds are obviously derived from (Z)-9-tricosene by oxidative metabolism (Blomquist et al., 1984). A microsomal P450 was shown to oxidize the alkene from either side at a distance of 9/10 carbons in-chain (Ahmad et al., 1987), but the structural requirements of the enzyme for its substrate are otherwise strict (Latli and Prestwich, 1991). This P450 activity is found in various tissues of the male and female, including the epidermis and fat body, but the highest specific activity is found in male antennae (Ahmad et al., 1987; Chapter 3.15). The C23 epoxide and ketone are absent internally in females, but accumulate on the surface of the cuticle (Mpuru et al., 2001), suggesting the localization of this P450 activity in epidermal cells of female flies. More generally, P450 enzymes are probably involved in the biosynthesis of many insect pheromones and allomones, e.g., epoxides of polyunsaturated hydrocarbons in arctiid moths, disparlure of the gypsy moth (Brattsten, 1979a), or monoterpenes

34 Insect Cytochrome P450

Figure 19 Biosynthesis of (Z )-9-tricosene and components of the housefly sex pheromone. P450hyd acts as a decarbonylase to produce (Z )-9-tricosene from an aldehyde precursor. Another P450 attacks (Z )-9-tricosene from either side to give the two components of muscalure.

in bark beetles (Brattsten, 1979a; White et al., 1979; Hunt and Smirle, 1988). In the honeybee, castespecific o and o-1 hydroxylations of fatty acids to mandibular pheromones (see Chapter 3.13) have all the characteristics of P450 reactions (Plettner et al., 1998). These P450s involved in pheromone biosynthesis are likely to be exquisitely specific. Evidence for pheromone catabolism by P450 enzymes is also accumulating. As shown for the metabolism of (Z)-9-tricosene in the housefly, a distinction between biosynthesis and catabolism can be purely semantic in the case of a biogenetic succession of chemicals that have different signaling

functions. Compound B that is a metabolite of compound A may have less or none of compound A’s activity, but may have its own specific biological activity. Nonetheless, pheromones as signal molecules need to be metabolically inactivated and this catabolism may occur in the antennae themselves. Several P450 mRNAs have been identified in insect antennae. In Drosophila, a partial P450 cDNA, along with a UDP-glycosyltransferase and a short chain dehydrogenase/reductase, were found by northern blot analysis to be preferentially expressed in the third antennal segments, with lower expression in legs (Wang et al., 1999). This P450 cDNA

Insect Cytochrome P450 35

Figure 20 P450-dependent N-demethylation and ring hydroxylation of the Phyllopertha diversa sex pheromone by antennal microsomes of male scarab beetles. See text and Wojtasek and Leal (1999) for details.

corresponds to Cyp6w1 for which many ESTs have been identified in a head cDNA library. In Mamestra brassicae, two P450 cDNAs were cloned from antennae, CYP4L4 and CYP4S4 (Maibeche-Coisne et al., 2002). Both genes are strongly expressed in the sensilla trichodea as shown by in situ hybridization. Whereas CYP4S4 expression is restricted to the antennae, CYP4L4 is also expressed in proboscis and legs. Five P450 ESTs representing three P450 genes were found in a small EST project using male Manduca sexta antennae (Robertson et al., 1999), which is further evidence that antennae may harbor several P450 enzymes. High levels of P450 reductase expression have been noted in Drosophila antennae (Hovemann et al., 1997), and adrenodoxin reductase is also expressed in antennae (Freeman et al., 1999). Antennal P450s are therefore an active enzyme system, complete with their redox partners. The physiological role of the P450s thus found in antennae is still formally unknown, but the presence of other enzymes generally associated with detoxification processes (Robertson et al., 1999; Wang et al., 1999) are indeed suggestive of a large category of odorant-metabolizing enzymes. In addition to the presence of P450 transcripts in antennae, there is at least one report where a P450 reaction has been characterized in antennae, but the CYP gene coding for the P450 catalyzing this reaction is still unknown. Antennal microsomes of the pale brown chafer, Phyllopertha diversa, metabolize the alkaloid sex pheromone by a P450 enzyme (Wojtasek and Leal, 1999). This enzyme is specifically produced in male antennae, and its activity is not detected in other scarab beetles or lepidopteran species (Figure 20). P450 enzymes may have found in insects a role as odorant degrading enzymes (ODE) along with esterases, aldehyde oxidases, glucosyl-transferases, etc. (review: Chapter 3.15). 4.1.4.1.5. Metabolism of fatty acids and related compounds P450-catalyzed fatty acid o-hydroxylation has been reported in insects (Feyereisen and

Durst, 1980; Ronis et al., 1988; Clarke et al., 1989; Cuany et al., 1990; Rose et al., 1991). Clofibrate selectively induces o-hydroxylation (and not o-1) of lauric acid by housefly and Drosophila microsomes, just as it induces the rat CYP4A1 o-hydroxylase (Clarke et al., 1989; Amichot et al., 1998). No clear peroxisome proliferator activated receptor (PPAR, Issemann and Green, 1990) gene ortholog but several paralogs are present in the Drosophila genome. In contrast to clofibrate, phenobarbital induces Drosophila o, o-1, and o-2 hydroxylations (Amichot et al., 1998), indicating the presence of several fatty acid hydroxylases. By homology to vertebrate CYP4 enzymes it is thought that cockroach CYP4C1 has a o-hydroxylase function as well, but evidence is lacking. CYP4C7, CYP6A1, and CYP12A1 lack lauric acid hydroxylase activity (Andersen et al., 1997; Guzov et al., 1998; Sutherland et al., 1998), but CYP6A8 has laurate o-1 hydroxylase activity (C. Helvig, personal communication). Although the role of vertebrate P450 enzymes in the metabolism of arachidonic acid and eicosanoids is well established (Capdevila et al., 2002), there is currently no indication for a similar function of P450 in insects. The biosynthesis of volicitin, an elicitor of plant volatile production found in the oral secretions of caterpillars occurs in the insect from the plantderived fatty acid linolenic acid (Pare et al., 1998). This biosynthesis involves C-17 hydroxylation and glutamine conjugation. It is likely that this hydroxylation will be shown to be catalyzed by a P450 enzyme. 4.1.4.1.6. Defensive compounds The tremendous variety of chemicals used for defense of insects against predation is well documented, but the enzymes involved in their de novo synthesis or in their transformation from ingested precursors are less studied. It is very likely that P450 enzymes may have found there a fertile ground for their chemical prowess. Just two examples are described. The biosynthesis of cyanogenic compounds found as defensive compounds in many insect species (Nahrstedt, 1988) may well involve P450 enzymes. For instance, the cyanogenic glucosides linamarin and lotaustralin found in Heliconius butterflies and Zygaena moths, are clearly derived from the amino acids valine and isoleucine. The pathway probably involves N-hydroxylation of the amino acids, further metabolism to the aldoximes and nitriles, and final C-hydroxylation before conjugation to the glucoside (Figure 21). The aldoximes and nitriles are efficiently incorporated in vivo (Davis and Nahrstedt, 1987; Holzkamp and Nahrstedt, 1994).

36 Insect Cytochrome P450

Another comparison of interest will be that of enzymes involved in the biosynthesis of insect defensive steroids with the well characterized steroid metabolizing enzymes of vertebrates. A number of aquatic Coleoptera (Dysticidae) and Hemiptera (Belastomatidae) synthesize a variety of steroids, mostly pregnanes (Scrimshaw and Kerfoot, 1987). With cholesterol as the presumed precursor in these carnivorous insects, one may envisage the evolution of an insect side-chain cleavage enzyme, of a C-21 hydroxylase and of a C17-C21 lyase that would catalyze reactions identical to those of CYP11A, CYP21, and CYP17. Some defensive steroids also have an aromatic A-ring, 7a, or 15a hydroxyl groups. A pregnene-3b, 20b-diol glucoside is synthesized from cholesterol in female pupae of M. sexta (Thompson et al., 1985). The role of this compound is unknown, but its synthesis strongly suggests the existence of a C20-C22 side chain cleavage enzyme in Lepidoptera as well. 4.1.4.2. Xenobiotic Metabolism: Activation and Inactivation

Figure 21 Biosynthesis of linamarin and lotaustralin from valine (R›H) and isoleucine (R›CH3), indicating possible sites of P450 metabolism. The efficiency of incorporation of intermediates in vivo is shown on the right. The homologous reactions catalyzed by two multifunctional Sorghum P450 enzymes converting tyrosine to dhurrin are shown on the left.

The pathway resembles that found in plants, where two multifunctional P450 enzymes are sufficient to convert the amino acid to the hydroxynitrile substrate of the conjugating enzyme (Figure 21). For dhurrin biosynthesis from tyrosine in sorghum, these are CYP79A1 and CYP71E1 (Kahn et al., 1997). Whereas the plant pathway appears to ‘‘channel’’ the substrates through the two P450s with little escape of the aldoxime intermediate, nothing is known of the number and functioning of the cyanogenic pathway enzymes used by Lepidoptera. Characterization of the P450 enzymes involved should allow a comparison of the plant and insect solutions to this biosynthetic challenge.

4.1.4.2.1. Natural products The metabolism of plant toxins by insects has been reviewed extensively (see, e.g., Brattsten, 1979b; Dowd et al., 1983; Ahmad, 1986; Ahmad et al., 1986; Mullin, 1986; Yu, 1986). Relatively a few studies have directly assessed the role of P450 enzymes in the metabolism of natural compounds by more than one or two criteria such as microsomal localization, NADPH and O2 dependence, and inhibition by piperonyl butoxide. In most cases, metabolism is associated with detoxification, e.g., the metabolism of xanthotoxin in Papilio polyxenes, S. frugiperda, and Depressaria pastinacella (Bull et al., 1986; Nitao, 1990), the metabolism of a-terthienyl in larvae of three lepidopteran species (Iyengar et al., 1990), and the metabolism of nicotine in Manduca sexta larvae (Snyder et al., 1993). Alpha- and beta-thujones are detoxified by P450 as evidenced by synergism of their toxicity by three P450 inhibitors in Drosophila and by the lower toxicity of six of their metabolites (Hold et al., 2001). Studies with flavone (Wheeler et al., 1993), monoterpenes (Harwood et al., 1990) and the alkaloid carnegine (Danielson et al., 1995) show clearly however that evidence for metabolism by P450 in vitro may not be sufficient to define an in vivo toxicological outcome. Natural products in the diet can act as inducers of P450 as well of other enzymes (e.g., glutathione S-transferases) and as a result of this induction (see below), the metabolism of the inducing compound or of coingested plant compounds can change dramatically over time (Brattsten et al., 1977).

Insect Cytochrome P450 37

Differences between acute and chronic toxicity are thus often the result of altered expression patterns (quantitative and qualitative) of P450 genes. This was demonstrated, for instance, in studies on Spodoptera larvae (Brattsten, 1983; Gunderson et al., 1986). The monoterpene pulegone and its metabolite menthofuran are more acutely toxic to S. eridania than to S. frugiperda, but the reverse is true for chronic toxicity. A study by Yu (1987) compared the metabolism of a large number of plant chemicals of different chemical classes by S. frugiperda and Anticarsia gemmatalis (velvetbean caterpillar) microsomes. Two indirect methods were used, on the one hand, the NADPH-dependent decrease in substrate and on the other hand, the substrate-induced NADPH oxidation. This metabolism is inhibited by piperonyl butoxide and by carbon monoxide, and induced by a number of chemicals, particularly indole-3carbinol, strongly suggesting P450 involvement. Such indirect methods are very useful as screening tools, as a first step towards a more thorough characterization of metabolism. However, they give no qualitative indication of the chemical fate of the substrate, nor quantitative indication of the levels of metabolism, as NADPH consumption is correlated to the coupling rate of the reaction, rather than to the rate of product formation (see Section 4.1.3.4.1). Clearly, insect P450 enzymes as a whole are capable of metabolizing a tremendous variety of naturally occurring chemicals, but the role of individual enzymes and their catalytic competence still needs a better description. Heterologously expressed P450 enzymes of the CYP6B subfamily from Papilio species (see Section 4.1.4.3 and Table 1) are well characterized for their ability to metabolize furanocoumarins (Hung et al., 1997; Li et al., 2003; Wen et al., 2003). They fit the description of enzymes with ‘‘broad and overlapping specificity’’ towards these compounds. Their range of catalytic competence is quite variable. For instance, CYP6B21 and CYPB25 metabolize the angular furanocoumarin angelicin at a similar rate (0.4–0.5 nmol/min/nmol P450), but whereas CYP6B21 also metabolizes 7-ethoxycoumarin at a similar rate (0.5 nmol/min/nmol P450), CYP6B25 does not have appreciable 7-dealkylation activity (Li et al., 2003). Natural products are not just an endless catalog of P450 substrates and inducers, but they also comprise a varied and complex set of inhibitors of P450 enzymes. These inhibitors range from ‘‘classical’’ reversible inhibitors to substrates that are activated to chemically reactive, cytotoxic forms (e.g., Neal and Wu, 1994).

4.1.4.2.2. Insecticides and other xenobiotics The metabolism of insecticides by P450 enzymes is very often a key factor in determining toxicity to insects and to nontarget species, but it can also represent a key step in the chain of events between contact, penetration, and interaction at the target site. The classical example is probably the metabolism of phosphorothioate insecticides. In many cases, the active ingredients of organophosphorus insecticides are phosphorothioate (P›S) compounds (a.k.a. phosphorothionates), whereas the molecule active at the acetylcholinesterase target site is the corresponding phosphate (P›O). It has long been recognized that the P›S to P›O conversion is a P450-dependent reaction. In the case of diazinon, this desulfuration has been studied for three heterologously expressed insect P450 enzymes (Dunkov et al., 1997; Guzov et al., 1998; Sabourault et al., 2001). All three P450s metabolized diazinon not just to diazoxon, the metabolite resulting from desulfuration, but also to a second metabolite resulting from oxidative ester cleavage. Similarly, antibodies to housefly CYP6D1 inhibit the microsomal desulfuration of chlorpyriphos as well as its oxidative ester cleavage (Hatano and Scott, 1993). The mechanism of P450-dependent desulfuration is believed to involve the initial attack of the P›S bond by an activated oxygen species of P450, leading to an unstable and therefore hypothetical phosphooxythiirane product (Figure 22). The collapse of this product can lead to two possible outcomes: (1) the replacement of sulfur by oxygen in the organophosphate product with the release of a reactive form of sulfur; and (2) the cleavage of the phosphate ester (or thioester) link with the substituent of highest electron-withdrawing properties, the ‘‘leaving group.’’ Outcome (1) can be viewed as ‘‘activation’’ because the P›S to P›O desulfuration produces an inhibitor of acetylcholinesterase often several orders of magnitude more potent than the P›S parent compound. However, the fate of this product of ‘‘activation’’ depends on the histological proximity to the target, sequestration, excretion, and further metabolism of the phosphate (by oxidative or hydrolytic enzymes). Kinetic evidence with the heterologously expressed CYP6A1, CYP6A2, and CYP12A1, as well as immunological evidence with CYP6D1 indicate that these P450 enzymes do not metabolize the P›O product of the parent P›S compound they metabolize. Outcome (2) or ‘‘dearylation’’ is without question a detoxification because the oxidative cleavage of the ‘‘leaving group’’ yields compounds unable to inhibit acetylcholinesterase, the dialkylphosphorothioate,

p0595

38 Insect Cytochrome P450

f0110

Figure 22 Metabolism of diazinon by cytochrome P450. Following an insertion of oxygen into the substrate, a reactive intermediate collapses (1) by desulfuration or (2) by cleavage of the ester linkage (dearylation). DEP: diethylphosphate, DEPT: diethylphosphorothioate, P-ol: 2-isopropopoxy-4-methyl-6-hydroxypyrimidine, [S]: reactive form of sulfur released during the reaction. Diazoxon can be further converted to DEP and P-ol by the same or another P450 in a subsequent reaction, or by a phosphotriester hydrolase. DEP may also be formed by spontaneous degradation of the initial product of diazinon monooxygenation. The ratio of outcomes 1 and 2 and the fate of the reactive sulfur depends on the P450 enzyme and on the type of OP substrate (see Table 4).

and/or dialkylphosphate. The ratio of outcome (1) and (2) appears P450-specific and substrate specific (Table 4) suggesting that the collapse of the unstable initial product of P450 attack is influenced by the active site environment. Theoretically, some P450 enzymes may very strongly favor outcome (1) or (2) or vice versa, thus qualifying as relatively ‘‘clean’’ activators or detoxifiers, but there is to date little direct evidence from the insect toxicological literature for such P450 enzymes (see however Oi et al., 1990). Furthermore, the sulfur released by the reaction can bind covalently either to neighboring proteins thus leading to cellular damage, or to the P450 protein itself (at least in vertebrate liver where this specific aspect has been studied, Kamataki and Neal, 1976). Parathion causes NADPH-dependent inhibition of methoxyresorufin O-demethylation activity (a P450Lpr-selective activity) in the housefly whereas chlorpyriphos does not (Scott et al., 2000). Therefore, it is not just the fate of the initial P450 metabolite of the P›S compound that depends on the P450 and the OP, but it is also the fate of the sulfur released that can vary.

Changes in the level of expression of P450 genes or P450 point mutations may be sufficient to change this delicate balance between activation and inactivation in vivo. For instance, fenitrothion resistance in the Akita-f strain of the housefly is related to an increase in oxidative ester cleavage over desulfuration measured in abdominal microsomes (Ugaki et al., 1985). In H. virescens, methyl parathion resistance in the NC-86 strain, which has an unchanged level of total P450, is related to a replacement of a set of P450 enzymes with high desulfuration activity by a set of P450 enzymes that metabolize less parathion, and do so with a lower desulfuration/oxidative ester cleavage ratio (Konno and Dauterman, 1989) (see Table 4). P450 enzymes that metabolize OPs can metabolize other insecticides as well and this sometimes leads to potentially useful interactions. Thus, enhanced detoxification of dicofol in spider mites can lead to enhanced chlorpyriphos activation, hence negative cross-resistance (Hatano et al., 1992). Similarly, permethrin resistance in horn flies is suppressible by piperonyl butoxide and negatively related to diazinon toxicity (Cilek et al., 1995). In H. armigera

p0605

Insect Cytochrome P450 39

t0020

Table 4 Desulfuration and oxidative ester cleavage of organophosphorus insecticides by P450 enzymes and microsomes

P450 enzyme

Ratio of OP desulfuration/oxidative ester cleavage

Human CYP2C19

8.5 d, 1.30 p, 0.14 c

Human CYP3A4 Human CYP2B6 Human liver microsomes Housefly CYP6A1 Drosophila CYP6A2 Housefly CYP12A1 Housefly CYP6D1 Housefly CSMA microsomes Housefly Akita-f a microsomes Heliothis virescens microsomes Heliothis virescens NC-86 a microsomes

3.0 d, 0.50 p, 0.66 c 0.7 d, 0.01 p, 3.38 c 0.29 d, 0.37 p, 0.57c 0.37 d 0.92 d 0.69 d 2.0 c 0.95 f 0.59 f 1.90 mp 1.32 mp

Reference

Kappers et al. (2001), Tang et al. (2001), Mutch et al. (2003) Kappers et al. (2001), Mutch et al. (2003) Kappers et al. (2001), Mutch et al. (2003) Kappers et al. (2001), Mutch et al. (2003) Sabourault et al. (2001) Dunkov et al. (1997) Guzov et al. (1998) Hatano and Scott (1993) Ugaki et al. (1985) Ugaki et al. (1985) Konno and Dauterman (1989) Konno and Dauterman (1989)

a Resistant strain. d, diazinon; p, ethyl parathion; c, chlorpyriphos; f, fenitrothion; mp, methyl parathion.

populations from West Africa, triazophos shows negative cross-resistance with pyrethroids, and in this case the synergism shown by the OP towards the pyrethroids appears to be due to an enhanced activation to the oxon form (Martin et al., 2003). These interactions were observed in vivo or with microsomes, but it is likely that they do reflect the properties of single P450 enzymes with broad substrate specificity rather than the fortuitous coordinate regulation of different P450 enzymes with distinct specificities. Organophosphorus compounds such as disulfoton and fenthion can also be activated by thioether oxidation (formation of sulfoxide and sulfone), but it is not clear whether these reactions are catalyzed in insects by a P450 or by a flavin monooxygenase (FMO). Further examples of oxidative bioactivation of organophosphorus compounds have been discussed (Drabek and Neumann, 1985). The toxicity of fipronil to house flies is increased sixfold by the synergist piperonyl butoxide, whereas the desulfinyl photodegradation product is not detoxified substantially by P450 (Hainzl and Casida, 1996; Hainzl et al., 1998). Conversion of fipronil to its sulfone appears to be catalyzed by a P450 enzyme in Ostrinia nubilalis (Durham et al., 2002) and in Diabrotica virgifera (Scharf et al., 2000). In the latter, the toxicity of fipronil sulfone is about the same as that of the parent compound, and piperonyl butoxide has only a marginal effect as synergist. In contrast, synergists antagonize the toxicity of fipronil in Blattella germanica, suggesting that oxidation to the sulfone represents an activation step in this species (Valles et al., 1997). The now banned cyclodiene insecticides aldrin, heptachlor, and isodrin are epoxidized by P450

enzymes to environmentally stable, toxic epoxides, dieldrin, heptachlor epoxide, and endrin (Brooks, 1979; Drabek and Neumann, 1985). Recombinant CYP6A1, CYP6A2, and CYP12A1 can catalyze these epoxidations (see Table 1). Examples of proinsecticide metabolism include the activation of chlorfenapyr by N-dealkylation (Black et al., 1994) and of diafenthiuron by S-oxidation (Kayser and Eilinger, 2001). In each case, the insect P450dependent activation is a key in the selective toxicity of these proinsecticides that target mitochondrial respiration. Recombinant housefly CYP6A1 catalyzes the activation of chlorfenapyr (V.M. Guzov, M. Kao, B.C. Black, and R. Feyereisen, unpublished data). In H. virescens, toxicity of chlorfenapyr is negatively correlated with cypermethrin toxicity (Pimprale et al., 1997). Genetic analysis indicates that a single factor is involved so the same P450 that activates chlorfenapyr may also detoxify cypermethrin in this species (Figure 23). A similar case of negative crossresistance of chlorfenapyr in a pyrethroid-resistant strain has been reported in the hornfly Haematobia irritans (Sheppard and Joyce, 1998). The metabolism of imidacloprid is also of interest in this respect. Although not extensively studied to date, there is evidence that piperonyl butoxide can synergize the toxicity of imidacloprid, but P450dependent metabolism can also lead to several bioactive metabolites in some insects. How these are further metabolized and how resistance can be caused by P450 attack on this molecule remains unclear (see however Section 4.1.4.5.5). In vivo synergism by piperonyl butoxide, a typical inhibitor of P450 enzymes (see Section 4.1.4.5.1), is often used to implicate a P450-mediated detoxification, and there are innumerable such examples in

40 Insect Cytochrome P450

Figure 23 Chlorfenapyr and cypermethrin metabolism. The same P450 in Heliothis virescens probably activates the pyrrole and inactivates the pyrethroid, resulting in negative cross-resistance.

the literature. The inference is much stronger when two unrelated synergists are used in vitro, and when metabolites of the pesticide are identified. For instance, pyriproxifen is hydroxylated by fat body and midgut microsomes of larval house flies to 40 -OHpyriproxyfen and 500 -OH-pyriproxyfen and these activities are inhibited by PB and TCPPE (Zhang et al., 1998). The study of xenobiotic metabolism by individual P450 enzymes expressed in heterologous systems has barely begun (Table 1). Whereas the CYP6 enzymes clearly comprise some enzymes with ‘‘broad and overlapping’’ substrate specificity, even closely related enzymes of this family can differ substantially in their catalytic competence. The task of predicting which xenobiotic or natural product will be metabolized by which type of P450 is currently not possible. 4.1.4.3. P450 and Host Plant Specialization

4.1.4.3.1. The Krieger hypothesis and beyond The interactions of plants and insects, and more specifically the role of plant chemistry on the specialization of phytophagous insects have generated a vast literature. ‘‘Secondary’’ plant substances are variously seen to regulate insect behavior and/or to serve as weapons in a coevolutionary ‘‘arms race’’ (Dethier, 1954; Fraenkel, 1959; Ehrlich and Raven, 1964; Jermy, 1984; Bernays and Graham, 1988). In chemical ecology alone, ‘‘no other area is quite so rife with theory’’ (Berenbaum, 1995). Many of the theories and some of the experiments implicitly or explicitly deal with the insect’s ability to metabolize plant secondary substances by P450 and other

enzymes. In the case of behavioral cues, we are far from understanding the true importance of P450 enzymes in the integration of chemosensory information, e.g., as ‘‘odorant degrading enzymes.’’ In the case of detoxification, however, the landmark paper of Krieger et al. (1971) can be seen as echoing the Fraenkel (1959) paper, by exposing the raison d’eˆtre of P450 enzymes. They stated that ‘‘higher activities of midgut microsomal oxidase enzymes in polyphagous than in monophagous species indicates that the natural function of these enzymes is to detoxify natural insecticides present in the larval food plants.’’ In that 1971 study, aldrin epoxidation was measured in gut homogenates of last instar larvae from 35 species of Lepidoptera. Polyphagous species had on average a 15 times higher activity than monophagous species. This trend was seen in sucking insects as well, with a 20-fold lower aldrin epoxidase activity in the oleander aphid Aphis nerii when compared to the potato aphid Myzus euphorbiae or to the green peach aphid M. persicae (Mullin, 1986). The former is a specialist feeder on two plant families, Asclepiadaceae and Apocyanaceae, whereas the latter two are generalists found on 30–72 plant families. The concept extended to other detoxification enzymes and was broadened to cover prey/predator, e.g., in mites where the predatory mite has a five times lower aldrin epoxidase activity than its herbivorous prey (Mullin et al., 1982). The toxicity of the natural phototoxin a-terthienyl is inversely propotional to the level of its metabolism in Lepidoptera and is related to diet breadth. Metabolism (4.0 nmol/min/nmol P450) is highest in O. nubilalis that feeds on numerous

Insect Cytochrome P450 41

phototoxic Asteraceae, lower in H. virescens that has a broad diet, including some Asteraceae that are nonphototoxic, and lowest in M. sexta, a specialist of Solanaceae (Iyengar et al., 1990). The conceptual framework of Krieger et al. has been challenged (Gould, 1984) and defended (Ahmad, 1986). An alternative view (Berenbaum et al., 1992) proposes that aldrin epoxidation represents ‘‘P450s with broad substrate specificity [that] are most abundant in insects that encounter a wide range of host plant metabolites.’’ A careful repetition of the Krieger experiments on lepidopteran larvae from 58 species of New South Wales failed to show significant differences in aldrin epoxidation between monophagous and polyphagous species (Rose, 1985). High activity in both monophagous and polyphagous species was invariably linked to the presence of monoterpenes in the host diet. The evidence presented in the sections below indicates that polyphagous and oligophagous species alike rely on the ability to draw on a great diversity of P450 genes, encoding a great diversity of specific and less specific enzymes and regulated by a great diversity of environmental sensing mechanisms – induction. The ability to induce P450 enzymes and deal with a wide range of toxic chemicals in the diet has been thought to present a ‘‘metabolic load’’ for polyphagous species, with specialists restricting their ‘‘detoxification energy’’ to one or a few harmful substrates (e.g., Whittaker and Feeny, 1971). However, careful studies in both oligophagous and polyphagous species have refuted this concept of metabolic load (e.g., Neal, 1987; Appel and Martin, 1992). 4.1.4.3.2. Host plant chemistry and herbivore P450 Cactophilic Drosophila species from the Sonoran desert are specialized to specific columnar cactus hosts by their dependency on unusual sterols (D. pachea) or by their unique ability to detoxify their host’s allelochemicals, notably isoquinoline alkaloids and triterpene glycosides (Frank and Fogleman, 1992; Fogleman et al., 1998). P450mediated detoxification was shown by the loss of larval viability in media that contained both allelochemicals and piperonyl butoxide, and by the induction of total P450 or alkaloid metabolism by the cactus allelochemicals or by phenobarbital (Frank and Fogleman, 1992; Fogleman et al., 1998). Several P450s of the CYP4, CYP6, CYP9, and CYP28 families are induced by cactus-derived isoquinoline alkaloids and by phenobarbital, but not by triterpene glycosides; only a CYP9 gene was induced by alkaloids and not by phenobarbital (Danielson et al., 1997, 1998; Fogleman et al., 1998). The capacity to detoxify isoquinoline alkaloids was not related to

DDT or propoxur tolerance, and while phenobarbital induced P450s capable of metabolizing the alkaloid carnegine in D. melanogaster, this was not sufficient to produce in vivo tolerance (Danielson et al., 1995). Selection of D. melanogaster with Saguaro alkaloids over 16 generations, however, led to P450-mediated resistance to the cactus alkaloids (Fogleman, 2000). These studies suggest the evolution of specific responses in the cactophilic species involving the recruitment of a phylogenetically unrelated subset of P450 genes in each instance of specialization of a fly species on its host cactus. The oligophagous tobacco hornworm (M. sexta) feeds essentially on Solanaceae and its adaptation to the high levels of insecticidal nicotine found in tobacco depends largely on metabolic detoxification, although other tolerance mechanisms may be contributing as well (Snyder and Glendinning, 1996). Hornworm larvae fed an nicotine-free artificial diet (naive insects) are rapidly poisoned by the ingestion of a nicotine-supplemented diet, but this diet is not deterrent. Poisoning is evidenced by convulsions and inhibition of feeding. The small amount of ingested nicotine induces its own metabolism, so that approximately 36 h later the larvae resume feeding normally, without further signs of poisoning. The inhibition of feeding and its resumption after nicotine exposure is directly related to P450 induction. Indeed, treatment with piperonyl butoxide, which itself has no effect on feeding, inhibits the increase in nicotine-diet consumption that occurs once nicotine metabolism has been induced (Snyder and Glendinning, 1996). Naive insects metabolize nicotine to nicotine 1-N-oxide at a low level, whereas nicotine-fed insects metabolize it further to cotinine-N-oxide at a higher level (Snyder et al., 1994). These reactions are catalyzed by one or more P450 enzymes (Snyder et al., 1993). The effects of nicotine on marker P450 activities are complex: the metabolism of three substrates is induced at low nicotine levels, seven are only induced at higher levels, and three are unaffected (Snyder et al., 1993). CYP4M1 and CYP4M3 are moderately induced in the midgut but not in the fat body, but CYP4M3 and CYP9A2 are not affected by nicotine (Snyder et al., 1995; Stevens et al., 2000). P450 induction has also been inferred in the polyphagous spider mite Tetranychus urticae, where the performance of a bean-adapted population on tomato was severely compromised by piperonyl butoxide (Agrawal et al., 2002). The P450 inhibitor did not reduce acceptance of tomato as a host, nor did it reduce the performance of the bean-adapted population on bean, strongly suggesting a postingestive induction of P450 as a mechanism of acclimation

42 Insect Cytochrome P450

to the novel host. In the polyphagous noctuid Spodoptera frugiperda, ingestion of indole 3carbinol increases once the continuous exposure to this toxic compound has induced P450 enzymes (Glendinning and Slansky, 1995). 4.1.4.3.3. Papilio species and furanocoumarins The adaptation of specialist herbivores to toxic components of their host plants is best documented in the genus Papilio. The black swallowtail, P. polyxenes, feeds on host plants from just two families, the Apiaceae (Umbelliferae) and the Rutaceae. These plants are phytochemically similar, particularly in their ability to synthesize furanocoumarins. Biogenetically derived from umbelliferone (7-hydroxycoumarin) the linear furanocoumarins (related to psoralen) and angular furanocoumarins (related to angelicin) are toxic to nonadapted herbivores (Berenbaum, 1990). This toxicity is enhanced by light as furanocoumarins are best known for their UV photoreactivity leading to adduct formation with macromolecules, particularly DNA. Papilio polyxenes has become a model in the study of adaptation to dietary furanocoumarins. Xanthotoxin, a linear furanocoumarin, induces its own metabolism in a dose-dependent fashion when added to the diet of P. polyxenes larvae (Cohen et al., 1989). This P450-dependent metabolism proceeds probably by an initial epoxidation of the furan ring followed by further oxidative attack and opening of the ring, leading to nontoxic hydroxylated carboxylic acids (Ivie et al., 1983; Bull et al., 1986). Inducible xanthotoxin metabolism is observed in all leaf-feeding stages of P. polyxenes, and is higher in early instars (Harrison et al., 2001). Xanthotoxin induces its own metabolism in the midgut, but also in the fat body and integument (Petersen et al., 2001). The metabolism of bergapten and sphondin is also induced by dietary xanthotoxin. Levels of total midgut microsomal P450s are unaffected by xanthotoxin, and photoactivation is not required for induction (Cohen et al., 1989). The metabolism of xanthotoxin is 10 times faster in P. polyxenes than in the nonadapted S. frugiperda (Bull et al., 1986). Papilio polyxenes microsomal P450s are also less sensitive to the inhibitory effects of xanthotoxin. NADPH-dependent metabolism of xanthotoxin leads to an uncharacterized reactive metabolite that can covalently bind P450 or neighboring macromolecules, i.e., xanthotoxin can act as a ‘‘suicide substrate’’ (Neal and Wu, 1994). This NADPH-dependent covalent labeling of microsomal proteins is seven times higher in M. sexta than in P. polyxenes. Inhibition of aldrin epoxidation and p-nitroanisole O-demethylation by xanthotoxin is also 6- and 300-fold higher, respectively, in M. sexta

than in P. polyxenes (Zumwalt and Neal, 1993). Myristicin, a methylene dioxyphenyl compound (see Section 4.1.3.4.4) found in the host plant parsnip is less inhibitory to P. polyxenes than to H. zea (Berenbaum and Neal, 1985; Neal and Berenbaum, 1989). A distinct protein band of 55 kDa appears in midgut microsomes of xanthotoxin-treated P. polyxenes larvae and its microsequencing led to the cloning of CYP6B1, a P450 shown to be inducible by xanthotoxin or parsnip (Cohen et al., 1992). Several variants of CYP6B1 have been cloned that presumably represent different alleles, v1, v2, and v3. The three variants differ from each other at 3, 6, or 9 amino acid positions (Cohen et al., 1992; Prapaipong et al., 1994). The CYP6B1 gene is selectively induced by linear furanocoumarins. Initial studies suggested that additional, related P450 transcripts were present in P. polyxenes and inducible by angular furanocoumarins (Hung et al., 1995b). Expression in the baculovirus system revealed that CYP6B1 v1 and v2 metabolize the linear furanocoumarins bergapten, xanthotoxin, isopimpinellin, and psoralen (Ma et al., 1994). Little metabolism of the angular furanocoumarin angelicin was observed in this early study but improvements in the heterologous expression system by coexpression of insect P450 reductase increased rates of metabolism sufficiently to confirm the role of CYP6B1 in the metabolism of angelicin as well (Wen et al., 2003). The furanocoumarins were metabolized in the improved expression system with the following preference: xanthotoxin > psoralen > angelicin. The latter is less efficiently metabolized in vivo (Li et al., 2003) and P. polyxenes is less adapted to it (Berenbaum and Feeny, 1981). A second P450 was cloned from P. polyxenes; it encodes CYP6B3 that is 88% identical to CYP6B1 (Hung et al., 1995a). CYP6B3 is expressed at lower basal levels than CYP6B1, but both CYP6B1 and CYP6B3 are inducible by xanthotoxin, sphondin, angelicin, and bergapten in the midgut. CYP6B3 responds more readily to the angular furanocoumarins than CYP6B1 (Hung et al., 1995a), and CYP6B1 is more inducible than CYP6B3 (Harrison et al., 2001). A later study showed that CYP6B1 is induced by xanthotoxin in the midgut, fat body, and integument, but CYP6B3 is induced by xanthotoxin only in the fat body (Petersen et al., 2001). The presence of CYP6B-like transcripts in species related to P. polyxenes was suggested early on by positive signals on northern blots with RNA from Papilio brevicauda and Papilio glaucus that were treated with xanthotoxin (Cohen et al., 1992). Papilio brevicauda is like P. polyxenes, a species

p0685

Insect Cytochrome P450 43

that feeds on furanocoumarin-containing Apiaceae, but Papilio glaucus is a generalist that encounters furanocoumarin-containing plants (e.g., hoptree, Ptelea trifoliata) only occasionally. Xanthotoxin, nevertheless, induces its own metabolism in all three species (Cohen et al., 1992). Papilio glaucus is highly polyphagous and is reported to feed on over 34 plant families, and therefore offers an interesting contrast to P. polyxenes. Esterase, glutathione S-transferase, and P450 activities are highly variable and dependent on the species of deciduous tree foliage that this species feeds on (Lindroth, 1989). Papilio glaucus has significant levels of linear and angular furanocoumarin metabolism, that are highly inducible by xanthotoxin (Hung et al., 1997). A series of nine CYP6B genes and some presumed allelic variants were cloned from P. glaucus (Hung et al., 1996, 1997; Li et al., 2001, 2002a). The first two genes CYP6B4 and CYP6B5 are products of a recent gene duplication event, and their promoter region is very similar (Hung et al., 1996). Six additional and closely related members of the CYP6B subfamily were cloned from Papilio canadensis, another generalist closely related to P. glaucus but not known to feed on plants containing furanocoumarins (Li et al., 2001, 2002a). Xanthotoxin induced CYP6B4-like and CYP6B17-like genes in both species, but the level of furanocoumarin metabolism was lower in P. canadensis (Li et al., 2001). This wide spectrum of CYP6B enzymes represents a wide range of activities towards furanocoumarin substrates. Whereas CYP6B4 of P. glaucus expressed in the baculovirus system efficiently metabolizes these compounds (Hung et al., 1997), CYP6B17 of P. glaucus, and CYP6B21 and CYP6B25 from P. canadiensis have a more modest catalytic capacity (Li et al., 2003). Papilio troilus, a relative of P. glaucus that specializes on Lauraceae that lack furanocoumarins, has undetectable basal or induced xanthotoxin metabolism (Cohen et al., 1992). The genus Papilio thus offers a complete range of situations: (1) specialists that deal efficiently with furanocoumarins by inducible expression of CYP6B genes; (2) generalists that also carry related CYP6B genes, but whose inducibility and metabolism are less efficient; and (3) nonadapted specialists that appear to have lost the inducible CYP6B panoply (Berenbaum et al., 1996; Berenbaum, 1999, 2002; Li et al., 2003). 4.1.4.3.4. Furanocoumarins and other insects The metabolism of furanocoumarins or the inducibility of P450 by these compounds is not restricted to Papilionidae. Xanthotoxin induces its own metabolism in the parsnip webworm, Depressaria

pastinacella (Nitao, 1989). This species belongs to the Oecophoridae, and is a specialist feeder on three genera of furanocoumarin-containing Apiaceae. It is highly tolerant to these compounds and metabolizes them not just by opening the furan ring, but in the case of sphondin, it is also capable of O-demethylation (Nitao et al., 2003). Although furanocoumarin metabolism is inducible, the basal (uninduced) activity is high (Nitao, 1989), and the response is a general one, with little discrimination of the type of furanocoumarin inducer or the type of furanocoumarin metabolized (Cianfrogna et al., 2002). The P450 enzymes involved in furanocoumarin metabolism by D. pastinacella are unknown. Low stringency northern hybridization failed to elicit a signal with a CYP6B1 probe (Cohen et al., 1992). A species that does not encounter furanocoumarins, the solanaceous oligophage M. sexta, responds to xanthotoxin by inducing CYP9A4 and CYP9A5 (Stevens et al., 2000). The generalist S. frugiperda also induces P450 as well as glutathione S-transferases in response to xanthotoxin (Yu, 1984; Kirby and Ottea, 1995). It has low basal P450mediated xanthotoxin metabolism, but this metabolism is inducible by a variety of compounds including terpenes and flavone (Yu, 1987). In the highly polyphagous H. zea, a similar situation is encountered. Xanthotoxin metabolism is low, but inducible by itself as well as by phenobarbital and a-cypermethrin (Li et al., 2000b). A number of CYP6B genes have been cloned from these Helicoverpa species. CYP6B8 of H. zea is very close in sequence to CYP6B7 from H. armigera, and it is inducible by xanthotoxin and phenobarbital (Li et al., 2000a). The high conservation of sequence in the SRS1 region suggests that the CYP6B enzymes of Helicoverpa are competent in furanocoumarin metabolism as indeed, these species occasionally encounter furanocoumarins in their diet. The CYP6B9/B27 and B8/B28 genes are pairs of recently duplicated genes (Li et al., 2002b). Their tissue and developmental pattern of expression is subtly different as is their pattern of induction by a variety of chemicals (Li et al., 2002c). 4.1.4.4. Host Plant, Induction and Pesticides

The adaptive plasticity conferred by the inducibility of P450 enzymes on different diets can have important consequences for insect control and the bionomics of pest insects. It is far from being just an ecological oddity or an interesting set of tales of insect natural history. It is well recognized that the same insect species fed different (host) plants will show differences in their response to pesticides

44 Insect Cytochrome P450

(Ahmad, 1986; Yu, 1986; Lindroth, 1991), and that these differences often reflect the induction of P450 enzymes, as well as of other enzymes, glutathione Stransferases, epoxide hydrolases, etc. The complexity of plant chemistry makes it difficult to account for the contribution of each individual chemical to this response and key components are often analyzed first (e.g., Moldenke et al., 1992). Similarly, the multiplicity of P450 genes and the range of P450 enzyme specificity makes it difficult to predict the outcome of exposure to a plant chemical. The toxicological importance of the plant diet on the herbivore’s P450 status (induction, inhibition) is well recognized in pharmacology where the joint use of chemical therapy and traditional herbs can have unpredicted outcomes (Zhou et al., 2003). Larvae of the European corn borer, Ostrinia nubilalis, fed leaves from corn varieties with increasing DIMBOA content and thus increasing levels of resistance to leaf damage had correspondingly increased levels of total midgut P450 and p-nitroanisole O-demethylation activity (Feng et al., 1992). These studies suggest that constitutive host plant resistance may affect the insect response to xenobiotics. In addition, the induction of host plant defense by insect damage may itself be a signal for induction of herbivore P450 enzymes, as shown in H. zea. The plant defense signal molecules jasmonate and salicylate induce CYP6B8, B9, B27, and B28 (Li et al., 2002d) in both fat body and midgut. The response to salicylate is relatively specific as p-hydroxybenzoate, but not methylparaben, also acts as an inducer. Treatment with 2-tridecanone, a toxic allelochemical from trichomes of wild tomato, protects H. zea larvae against carbaryl toxicity (Kennedy, 1984) and H. virescens larvae against diazinon toxicity (Riskallah et al., 1986b). In H. virescens larvae, the compound caused both qualitative and quantitative changes in P450 spectral properties (Riskallah et al., 1986a), an induction confirmed by its effect on specific P450 genes in the gut of M. sexta larvae (Snyder et al., 1995; Stevens et al., 2000). Larvae of H. virescens with a genetic resistance to 2-tridecanone have increased P450 levels and P450 marker activities (benzphetamine demethylation, benzo[a]pyrene hydroxylation, phorate sulfoxidation), and these can be further increased by feeding 2-tridecanone (Rose et al., 1991). A laboratory population of H. zea can rapidly display increased tolerance to a-cypermethrin by selection of an increased P450 detoxification ability with a high dose of dietary xanthotoxin (Li et al., 2000b). Beyond the host plant, it is the whole biotic and chemical environment that determines the response

of an insect to pesticide exposure. Herbicides and insecticide solvents can serve as inducers (Brattsten and Wilkinson, 1977; Kao et al., 1995; Miota et al., 2000). Aquatic larvae are exposed to natural or anthropogenic compounds that alter their P450 detoxification profile (David et al., 2000, 2002; Suwanchaichinda and Brattsten, 2002). Virus infection affects P450 levels (Brattsten, 1987), and the expression of several P450 genes is affected during the immune response (see Section 4.1.5.1.2). 4.1.4.5. Insecticide Resistance

4.1.4.5.1. Phenotype, genotype, and causal relationships Insecticide resistance is achieved in a selected strain or population by: (1) an alteration of the target site; (2) an alteration of the effective dose of insecticide that reaches the target; or (3) a combination of the two. The resistance phenotypes have long been analyzed according to these useful biochemical and physiological criteria. At the molecular genetic level, several classes of mutations can account for these phenotypes (Taylor and Feyereisen, 1996) and a causal relationship between a discrete mutation and resistance has been clearly established for several cases of target site resistance (ffrenchConstant et al., 1999). The molecular mutations responsible for P450-mediated insecticide resistance are only beginning to be explored. In contrast to CYP51, which is a target for a major class of fungicides, no insect P450 has been recognized as a primary target for a commercial insecticide. Thus, biochemical changes in P450 structure or activity can lead to changes in insecticide sequestration, activation, or inactivation, so that all the classes of molecular mutations (structural, up- or downregulation, see Taylor and Feyereisen, 1996) can be theoretically involved in P450-mediated resistance. When the number of P450 genes is taken into account, it is little wonder that P450 enzymes are so often involved in insecticide resistance, and that it has been so difficult finding, and establishing, the role of resistance mutations for P450 genes. Traditionally, the first line of evidence for a role of a P450 enzyme in resistance has been the use of an insecticide synergist (e.g., piperonyl butoxide), a suppression or decrease in the level of resistance by treatment with the synergist being diagnostic. In cases too many to list here, this initial and indirect evidence is probably correct, however there are cases where piperonyl butoxide synergism has not been explained by increased detoxification (Kennaugh et al., 1993). Piperonyl butoxide may also be a poor inhibitor of the P450(s) responsible for resistance, so that the use of a second, unrelated synergist may be warranted (Brown et al., 1996; Zhang et al.,

Insect Cytochrome P450 45

1997). In addition, the synergist as P450 inhibitor can decrease the activation of a proinsecticide, so that lack of resistance suppression can be misleading. Chlorpyriphos resistance in D. melanogaster from vineyards in Israel maps to the right arm of chromosome 2 (see Section 4.1.4.5.5) and is enhanced by piperonyl butoxide rather than suppressed (Ringo et al., 1995). An independent and additional line of evidence is the measurement of total P450 levels or metabolism of selected model substrates. An increase in either or both being viewed as diagnostic. Again, such evidence is tantalizing but indirect, and the absence of change uninformative. The validation of a model substrate for resistance studies requires substantial knowledge about the P450(s) involved, and is therefore best assessed a posteriori. An increase in the metabolism of the insecticide itself in the resistant strain is more conclusive. For instance, permethrin metabolism to 40 -hydroxypermethrin was higher in microsomes from Culex quinquefasciatus larvae that are highly resistant to permethrin (Kasai et al., 1998b) than in their susceptible counterparts. Total P450 and cytochrome b5 levels were 2.5 times higher in the resistant strain. Both permethrin toxicity and metabolism were inhibited by two unrelated synergists, TCPPE and piperonyl butoxide. A similarly convincing approach was taken to show P450 involvement in the resistance of housefly larvae of the YPPF strain to pyriproxifen. Gut and fat body microsomes were shown to metabolize the IGR to 40 -OH-pyriproxyfen and 500 -OH-pyriproxyfen at higher rates than microsomes of the susceptible strains and this metabolism was synergist-suppressible (Zhang et al., 1998). The major, dominant resistance factor was linked to chromosome 2 in that strain (Zhang et al., 1997). Increased levels of transcripts for one or more P450 genes in insecticide-resistant strains has now been reported in many cases (see Table 5). This suggests that overexpression of one or more P450 genes is a common phenomenon of metabolic resistance but does not by itself establish a causal relationship with resistance. In some cases, the increased mRNA levels have been related to increased transcription (Liu and Scott, 1998), or increased protein levels (Liu and Scott, 1998; Sabourault et al., 2001). Genetic linkage between increased mRNA or protein levels for a particular P450 and resistance has been obtained to the chromosome level (CYP6A1, Cyp6a2, Cyp6a8, CYP6D1, CYP9A1, CYP12A1: Carin˜ o et al., 1994; Liu and Scott, 1996; Rose et al., 1997; Guzov et al., 1998; Maitra et al., 2000), and closer to marker genes (Cyp6g1, CYP6A1: Daborn

et al., 2001; Sabourault et al., 2001). Linkage is just the first step in establishing a causal link between a P450 gene and resistance. The following is a discussion of specific cases of P450 genes associated with insecticide resistance that have been studied in greater detail. Evidence for mutations causing constitutive overexpression in cis and trans, as well as an example of point mutations in a P450 coding sequence are currently available. The variety of mechanisms, even in a single species in response to the same insecticide, is striking. The paucity of available data on the molecular definition of the resistant genotype and on its causal relationship to resistance is also striking when compared to the wealth of data on target site resistance (ffrench-Constant et al., 1999). 4.1.4.5.2. CYP6A1 and diazinon resistance in the housefly Rutgers strain CYP6A1 was the first insect P450 cDNA to be cloned, and the gene was shown to be phenobarbital-inducible and constitutively overexpressed in the multiresistant Rutgers strain (Feyereisen et al., 1989). A survey of 15 housefly strains (Carin˜ o et al., 1992) showed that CYP6A1 is constitutively overexpressed at various degrees in eight resistant strains, but not in all resistant strains, notably R-Fc known to possess a P450-based resistance mechanism. Thus, the first survey with a P450 molecular probe confirmed the results of the first survey of housefly strains with marker P450 activities (aldrin epoxidation and naphthalene hydroxylation; Schonbrod et al., 1968): there is no simple relationship between resistance and a molecular marker, here the level of expression of a single P450 gene. That different P450 genes would be involved in different cases of insecticide resistance was a sobering observation (Carin˜ o et al., 1992), even before the total number of P450 genes in an insect genome was known. The constitutive overexpression of CYP6A1 was observed in larvae and in adults of both sexes. Overexpression was shown in both developmental stages to be linked to a semidominant factor on chromosome 2 (Carin˜ o et al., 1994), but the CYP6A1 gene was mapped to chromosome 5 (Cohen et al., 1994). The gene copy number being identical between Rutgers and a standard susceptible strain (sbo), gene amplification could not be invoked to explain overexpression (Carin˜ o et al., 1994), and the existence of a chromosome 2 trans-acting factor(s) differentially regulating CYP6A1 expression in the Rutgers and sbo strains was implied. Competitive ELISA using purified recombinant CYP6A1 protein as standard showed that the elevated mRNA levels were indeed translated into elevated protein levels (Sabourault

46 Insect Cytochrome P450

t0025

Table 5 P450 overexpression in insecticide-resistant strains P450 overexpressed Musca domestica CYP6A1

CYP6A1a CYP6D1

P450Lpr/CYP6D1a CYP6D1/CYP6D1a CYP6D1 CYP6D3 CYP12A1 Drosophila melanogaster Cyp4e2 Cyp6a2

CYP6A2a Cyp6a8

Cyp6g1

Cyp12d1/2 Drosophila simulans CYP6G1b Heliothis virescens CYP9A1 Helicoverpa armigera CYP4G8 CYP6B7 Lygus lineolaris CYP6X1 Anopheles gambiae CYP6Z1 Culex quinquefasciatus CYP6F1 Culex pipiens pallens CYP4H21, H22, H23 CYP4J4, CYP4J6 Diabrotica virgifera CYP4 Blattella germanica

Strain

Resistance pattern

Reference

Rutgers and other strains Rutgers LPR LPR NG98, Georgia YPER LPR Rutgers

OP, carbamates, IGR

Feyereisen et al. (1989), Carin˜o et al. (1992) Sabourault et al. (2001) Liu and Scott (1996) Liu and Scott (1996) Kasai and Scott (2000) Shono et al. (2002) Kasai and Scott (2001b) Guzov et al. (1998)

RaleighDDT 91R RaleighDDT MHIII-D23 Several strains 91R MHIII-D23 Wis-1lab Hikone R and 20 strains WC2 EMS1 Wisconsin-1, 91-R Wisconsin-1, 91-R

b

Pyrethroids Pyrethroids

DDT DDT DDT Malathion DDT, pyrethroids

DDT DDT Lufenuron, propoxur Imidacloprid DDT DDT

Amichot et al. (1994) Waters et al. (1992) Amichot et al. (1994) Maitra et al. (2000) Bride et al. (1997) Maitra et al. (1996) Maitra et al. (2000) Le Goff et al. (2003) Daborn et al. (2001, 2002) Daborn et al. (2002) Daborn et al. (2002) Brandt et al. (2002) Brandt et al. (2002), Le Goff et al. (2003)

OV1

DDT, imidacloprid, malathion

Le Goff et al. (2003)

Macon Ridge

Thiodicarb

Rose et al. (1997)

Pyrethroids Pyrethroids

Pittendrigh et al. (1997) Ranasinghe and Hobbs (1998)

Permethrin

Zhu and Snodgrass (2003)

PSP

Pyrethroids

Nikou et al. (2003)

JPal-per

Pyrethroids

Kasai et al. (2000)

RR

Deltamethrin

Shen et al. (2003)

Me-parathion, carbaryl

Scharf et al. (2001)

Chlorpyriphos

Scharf et al. (1999)

P450MAa a

Pyrethroids

P450 protein level increased. Presumed CYP6G1 ortholog of D. simulans.

et al., 2001; see Table 6 for comparison of Rutgers and sbo). A comparison of the coding sequence of CYP6A1 between Rutgers and two susceptible strains showed no (sbo) or little (aabys) sequence variation. Five amino acid changes were noted in aabys, two at the far N-terminus and three at the far C-terminus, well outside the regions (SRS) thought to influence enzyme activity (Cohen et al., 1994). The lack of CYP6A1 sequence difference between

Rutgers and sbo indicated that if CYP6A1 was implicated in diazinon resistance in the Rutgers strain, it was through elevation of enzyme activity alone. Reconstitution of recombinant CYP6A1 expressed in E. coli with its redox partners (Sabourault et al., 2001) provided the conclusive evidence for its role in diazinon resistance, as CYP6A1 metabolizes the insecticide with a high turnover (18.7 pmol/pmol CYP6A1/min), and a

Insect Cytochrome P450 47

Table 6 Comparison of a susceptible and a resistant strain of the housefly

Fly strain

Diazinon contact toxicity: LC50 (mg/pint jar) Resistance ratio Diazinon topical toxicity: LD50 (mg/fly) Resistance ratio P450 level (nmol/mg protein) Aldrin epoxidation (pmol/min/pmol P450) CYP6A1 mRNA relative level CYP12A1 mRNA relative level CYP6A1 protein level (fmol/abdomen) Diazinon metabolized by CYP6A1 Oxidative cleavage (pmol/min/fly) Desulfuration to oxon (pmol/min/fly) OP oxon metabolized by mutant aliesterase (pmol/min/fly) NADPH-dep. diazinon metabolism by microsomes (pmol/min/abdomen) a

p0760

sbo (S )

Rutgers (R)

4.4

167.8

1 0.059a 1 0.14 4.4a

37.8 7.1 120 0.29 15.6

1 1 36 0.5 0.2 0.0 2.9a

27.5 15 565 7.7 2.9 2.2–2.5 15.5

NAIDM susceptible strain.

favorable ratio (2.7) between oxidative ester cleavage and desulfuration (see Section 4.1.4.2.2 and Table 4). The nature of the chromosome 2 trans-acting factor and of the mutation leading to resistance in the Rutgers strain has long remained enigmatic despite considerable circumstantial evidence for a major resistance factor on chromosome 2 (Plapp, 1984). Diazinon resistance and high CYP6A1 protein levels could not be separated by recombination in the short distance between the ar and car genes (3.3– 12.4 cM). This region carries an ali-esterase gene (MdaE7) implicated as its Lucilia cuprina ortholog in organophosphorus insecticide resistance (Newcomb et al., 1997; Claudianos et al., 1999). A Gly137 to Asp mutation in this ali-esterase abolishes carboxylesterase activity towards model compounds such as methylthiobutyrate, and confers a measurable phosphotriester hydrolase activity towards an organophosphate (‘‘P›O’’), chlorfenvinphos. Chromosome 2 of the Rutgers strain carries this Gly137 to Asp mutation, and low CYP6A1 protein levels are correlated with the presence of at least one wild-type (Gly137) allele of MdaE7. Recombination in the ar-car region could not dissociate diazinon susceptibility, low CYP6A1 protein level, and the presence of a Gly137 allele of the aliesterase (Sabourault et al., 2001). It was therefore hypothesized that the wild-type ali-esterase metabolizes an (unknown) endogenous substrate into a negative regulator of CYP6A1 transcription. Removal of this regulator (by loss-of-function of

the ali-esterase) would increase CYP6A1 production and, hence, diazinon metabolism. Nature seems to have found the optimal loss-of-function mutation (Gly137 to Asp) as the Rutgers haplotype has swept through global populations of the housefly (C. Claudianos, J. Brownlie, V. Taskin, M. Kence, R.J. Russell, J.G. Oakeshott, personal communication). The mutant ali-esterase probably helps clearing the activated form (P›O) of the insecticide (Sabourault et al., 2001). The negative regulation by the Gly137 allele and the diazinon resistanceenhancing effect of the Asp137 allele may explain the incomplete dominance of the diazinon resistance trait. Housefly strains that are susceptible or that are not known/shown to overexpress CYP6A1 predictably carry at least one Gly137 allele (Scott and Zhang, 2003). The LPR strain that overexpresses CYP6A1 (Carin˜ o et al., 1992) and has increased OP metabolism (Hatano and Scott, 1993), as well as another resistant strain (NG98) carry other alleles (Scott and Zhang, 2003) that have impaired ali-esterase activity, Trp251 to Ser or Leu (Campbell et al., 1998; Claudianos et al., 2001). Corroborating, but indirect evidence for the hypothesis is the predicted pleiotropic effect of a trans-acting regulator. Constitutive overexpression of CYP12A1 whose product metabolizes diazinon as well (Guzov et al., 1998) and GST-1 in the Rutgers strain are also controlled by a chromosome 2 factor, possibly the same as the one controlling CYP6A1 expression. Although alternative hypotheses can be advanced, such as the fortuitous genetic closeness between the ali-esterase and a factor that increases the level of CYP6A1, a diazinon metabolizing enzyme, such hypotheses do not have the benefit of elegance. The mutant ali-esterase cannot account for the carbamate and JHA resistance so is not the sole factor of resistance seen in many housefly strains. Diazinon-resistant L. cuprina have provided evidence for Oppenoorth’s mutant ali-esterase hypothesis (Newcomb et al., 1997), but the role of P450 in OP resistance cannot be ignored. Indeed, the Q strain of the sheep blowfly is more resistant to parathion than to paraoxon (Hughes and Devonshire, 1982) and indirect evidence for a P450 involvement in L. cuprina diazinon resistance has also been presented (Kotze, 1995; Kotze and Sales, 1995). 4.1.4.5.3. Cyp6a2 in Drosophila: overexpression and mutant enzyme Insecticide-resistant strains of D. melanogaster have been studied at the molecular genetic level and the Cyp6a2 gene has been implicated in the metabolic resistance of several of

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48 Insect Cytochrome P450

them. Cyp6a2 is constitutively overexpressed in the 91R strain that is resistant to malathion and DDT by a factor of 20–30 relative to the susceptible 91C strain (Waters et al., 1992). DDT resistance maps to 56 cM on the left arm of chromosome 2 in the 91R strain (Dapkus, 1992), which is at or near the chromosomal location of Cyp6a2 (43A1-2). Initially, the presence of a 96 bp insertion in the 30 UTR of the gene was proposed to confer a low level of expression to the Cyp6a2 gene (Waters et al., 1992), but this insertion (or rather, the lack of it) was neither correlated with DDT resistance (Delpuech et al., 1993) nor confirmed to be linked to overexpression in resistant strains (Dombrowski et al., 1998). Cyp6a8 is also constitutively overexpressed in the 91R strain (Maitra et al., 1996) and the expression of both Cyp6a2 and Cyp6a8 is repressed in 91R/ 91C hybrids (Maitra et al., 1996; Dombrowski et al., 1998; Maitra et al., 2000). Flies (rosy506, insecticide susceptible and with constitutively low expression of Cyp6a2) were transformed with a P-element carrying the Cyp6a2 gene of the 91R strain driver by its own promoter. These flies express the transgene at higher levels than the endogenous Cyp6a2 but at lower levels than in their native 91R background (Dombrowski et al., 1998). The expression of both Cyp6a2 and Cyp6a8 is also constitutively higher in the MHIII-D23 strain initially selected for malathion resistance (Maitra et al., 2000). Genetic crosses and chromosome substitution experiments conclusively showed that the expression of both genes (located on the 2R chromosome) is repressed by factors on the third chromosome of the insecticide-susceptible 91C and rosy506 strains. In contrast, the third chromosome of the MHIII-D23 and 91R strains carries a loss-offunction mutation for this negative trans regulator, allowing the constitutive overexpression of the two genes (Maitra et al., 2000). Further careful examination of the promoter activity of the Cyp6a8 gene by fusion with a luciferase reporter gene in transgenic flies identified a11/761 bp region of the Cyp6a8 gene of the 91R strain that was sufficient to respond to the negative regulation by the rosy506 (wild-type) trans acting factor (Maitra et al., 2002). CYP6A2, similar to CYP6A1, appears to have a broad substrate specificity (see Table 1). Whether CYP6A2 of the 91R strain is capable of metabolizing DDT is unknown, as its sequence differs from that of the baculovirus produced CYP6A2 from the iso-1 strain, which does not metabolize DDT, and from that of the Raleigh-DDT strain, which does (see below). Nonetheless, the studies with the 91R and MHIII-D23 strains are clear indications for loss-of-function mutations in gene(s) encoding

negative regulators of P450 gene expression on chromosome 3. Genetic analyses of malathion resistance and of P450 expression (electrophoretic bands and marker activities) (Hallstrom, 1985; Hallstrom and Blanck, 1985; Houpt et al., 1988; Waters and Nix, 1988) have pointed to one or more loci between 51 and 61 cM on the right arm of chromosome 3, and it is interesting that this chromosome arm is thought to be orthologous to chromosome 2 of the housefly (Weller and Foster, 1993). The Drosophila ortholog of the MdaE7 gene (Est23) is located at 84D9. Resistance to DDT in the RaleighDDT strain offers a different picture. This strain has very high DDT resistance (>10 000-fold; Cuany et al., 1990). Its piperonyl butoxide-suppressible resistance is polyfactorial but the major, dominant resistance factor maps to 55 cM on the second chromosome as in the 91R strain. At least two P450 genes, Cyp6a2 and Cyp4e2, are constitutively overexpressed in this strain (Amichot et al., 1994). The genetic localization of resistance matches the locus of Cyp6a2 (A. Brun-Barale, S. Tares, J.M. Bride, A. Cuary, J.B. Berge´ , M. Amichot, personal communication), and the RaleighDDT allele was sequenced. Three point mutations, Arg335 to Ser, Leu336 to Val, and Val476 to Leu, were found (Berge´ et al., 1998). Overexpression was separated from the point mutations by repeated backcrossing to a marked susceptible strain and DDT selection. The resulting strain called 152 retained the mutant Cyp6a2 gene and high monofactorial resistance to DDT (>1000-fold), which also maps to 54.4 cM. Both RaleighDDT and 152 flies are characterized by elevated in vitro DDT metabolism, as well as elevated ethoxycoumarin and ethoxyresorufin O-deethylase activities. Strain 152 lost the constitutive expression of Cyp6a2, which was therefore caused by an unlinked trans-acting factor (A. BrunBarale, S. Tares, J.M. Bride, A. Cuary, J.B. Berge´ , M. Amichot, personal communication). The wild-type CYP6A2 and five engineered versions (each individual mutation, a double mutant carrying the first two and a triple mutant corresponding to the RaleighDDT allele) were expressed in E. coli and assayed for activity. The mutant enzymes were all characterized by a decreased stability when compared to the wild-type enzyme. The enzyme production in E.coli was significantly lower and the ratio of holoenzyme produced (measured by the FeII–CO complex) to apoenzyme produced (measured with an antiCYP6A2 antibody) was also lower. However, the triple mutant was uniquely capable of metabolizing DDT to DDA, DDD, and dicofol at rates that were

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p0780

Insect Cytochrome P450 49

9–13 times higher than the wild-type enzyme whose DDT metabolism was barely measurable. In addition to the triple mutant, only the Arg335 to Ser single mutant had the capacity to hydroxylate DDT to dicofol at a rate significantly different from wildtype. In contrast to DDT metabolism, the 16b hydroxylation of testosterone was not affected in the various single and multiple mutants. Thus, the three mutations found in Cyp6a2 of the RaleighDDT (and its derivative 152) strain collectively confer DDT-metabolizing ability to the mutant CYP6A2 enzyme. Genetic localization of DDT resistance to the Cyp6a2 gene locus is therefore explained. When the point mutations of the 152 strain are combined with overexpression as in the parent RaleighDDT strain, the very high level of resistance can be rationalized. This is the first example of point mutations in an insect P450 enzyme that contribute to insecticide resistance.

p0785

4.1.4.5.4. CYP6D1, the housefly LPR strain, and pyrethroid resistance The LPR strain of the housefly is highly resistant to pyrethroids (see Chapter 6.1) with a phenoxybenzyl moiety. This permethrin-selected strain has several resistance mechanisms, with P450-based detoxification as a major contributor. An abundant form of P450 (P450Lpr) was purified from abdomens of adult LPR flies, and immunological data indicated that P450Lpr represents 67% of the P450 in microsomes from LPR flies, a tenfold (Wheelock and Scott, 1990) increase over the reference-susceptible strain. Peptide sequences from the purified protein allowed PCR amplification and cloning of the P450Lpr gene, CYP6D1 (Tomita and Scott, 1995). This gene is located on chromosome 1 of the housefly (Liu et al., 1995), and is constitutively overexpressed by about tenfold in the LPR strain. This overexpression is not caused by gene amplification, but by increased transcription. It has been claimed that increased transcription of CYP6D1 causes insecticide resistance (Liu and Scott, 1998), but transgenic expression of CYP6D1 in Drosophila (Korytko et al., 2000a) has not been reported to confer resistance, and CYP6D1 produced in yeast (Smith and Scott, 1997) has not been reported to metabolize pyrethroids. Instead, the evidence for the role of CYP6D1 in pyrethroid resistance is based on the inhibition of microsomal deltamethrin and cypermethrin metabolism by anti-P450Lpr antibodies (Wheelock and Scott, 1992b; Korytko and Scott, 1998). The metabolism of pyrethroids by CYP6D1 has been studied in its microsomal environment of the LPR strain. Deltamethrin is metabolized (200 A domains. Apparently, the enzyme is tethered to the cuticle by the ChBD, which anchors the catalytic domain to the insoluble substrate and localizes the hydrolysis of chitin to an area with a radius of several hundred angstroms. The use of such a tethered enzyme would help to prevent diffusion of the soluble enzyme from the insoluble polysaccharide. In the case of Tenebrio chitinase, which consists of five catalytic, five linker, and six chitin-binding domains (Royer et al., 2002), one could envision a situation where a much wider area of the chitin– protein matrix undergoes intensive degradation by a much larger tethered enzyme. 4.3.4.6. Mechanism of Catalysis

Insect chitinases are members of family 18 of the glycosylhydrolases (CAZY, 2004), which generally utilize a mechanism where retention of the anomeric configuration of the sugar donor occurs via a

substrate-assisted catalysis, rather than a mechanism similar to lysozyme, which involves a proton donor and an electrostatic stabilizer (Fukamizo, 2000). However, a recent kinetic study of bacterial family 18 chitinases demonstrated that substrates lacking the N-acetyl group and thus incapable of anchimeric assistance were nevertheless hydrolyzed, suggesting that the reaction mechanism of family 18 chitinases cannot be fully explained by the substrate-assisted catalysis model (Honda et al., 2003). Therefore, additional studies are still required to understand fully the reaction mechanism of family 18 chitinases. The interaction of insect chitinases with insoluble chitin in the exoskeleton and PM is rather complex and believed to be a dynamic process that involves adsorption via a substrate-binding domain, hydrolysis, desorption, and repositioning of the catalytic domain on the surface of the substrate. This degradative process apparently requires a coordinated action of multiple domains by a mechanism that is not well understood. In addition to the catalytic events, the mechanism of binding of the enzyme onto the heterogeneous surface of native chitin is poorly characterized. Hydrolysis of chitin to GlcNAc is accomplished by a binary enzyme system composed of a chitinase and a b-N-acetylglucosaminidase (Fukamizo and Kramer, 1985;

Figure 5 Ribbon (left) and space-filling (right) model structures of Manduca sexta chitinase with the catalytic and chitin-binding domains shown in complexes with chitin oligosaccharides (yellow). In the ribbon representation, the polypeptide chain is colorcoded, beginning with blue at the N-terminus and proceeding through the rainbow to red at the C-terminus. The catalytic domain structure (top) was modeled using the program SOD (Kleywegt et al., 2001) with human chitotriosidase (PDB entry code 1LG1) (Fusetti et al., 2002) serving as the template. The chitin-binding domain (bottom) was similarly obtained using tachychitin (PDB entry 1DQC) (Suetake et al., 2000) as the template. The linker region is shown as a random coil as predicted by secondary structure prediction software and supported by circular dichroism data. The oligosaccharides are shown as stick models (left) and spacefilling models (right). Substrate binding to the catalytic domain was modeled using the available structures of complexes from glycosyl hydrolase family 18, while binding to the chitin-binding domain was modeled based on sequence conservation within the subfamily. M. sexta chitinase is a glycoprotein that is glycosylated especially in the linker region; however, no carbohydrate is shown in the model. (The model was constructed by Wimal Ubhayasekera and Dr. Sherry Mowbray, Swedish University of Agricultural Sciences, Uppsala.)

Chitin Metabolism in Insects

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Filho et al., 2002). The former enzyme hydrolyzes the insoluble polymer into soluble oligosaccharides, whereas the latter further degrades the oligomers to the monomer from the nonreducing end. Mechanistically, chitinases of family 18 hydrolyze chitin with retention of the anomeric configuration at the cleavage site, involving a double-displacement mechanism where a substrate-assisted catalysis occurs (Tomme et al., 1995; Henrissat, 1999; Zechel and Withers, 2000; Brameld et al., 2002). B. mori chitinase utilizes a retaining mechanism, yielding products that retain the b-anomeric configuration (Abdel-Banat et al., 1999). All of the enzymes of this family are inhibited by allosamidin, a transition state analog inhibitor which apparently is diagnostic for enzymes that utilize the retaining mechanism (Koga et al., 1987; Bortone et al., 2002; Brameld et al., 2002; Lu et al., 2002). Analysis of the products from the hydrolysis of chitin oligosaccharides by the family 18 chitinase from S. marcescens revealed variable subsite binding preferences, anomeric selectivity, and the importance of individual binding sites for the processing of short oligosaccharides compared to the cumulative recognition and processive hydrolysis mechanism used to digest the polysaccharide (Aronson et al., 2003). Polysaccharide-hydrolyzing enzymes are known to exhibit nonideal kinetic behavior because they often are susceptible to inhibition by both substrates and products (Va¨ ljama¨e et al., 2001). All insect chitinases examined were found to be susceptible to inhibition by oligosaccharide substrates but to varying extents (Fukamizo and Kramer, 1985; Fukamizo et al., 1995; Fukamizo, 2000). Apparently, the oligosaccharide substrate molecules can bind to these enzymes in such a manner that none of the target bonds is properly exposed to the functional groups of catalytic amino acids or the substrate may bind in only noncatalytic subsites of the larger active site, forming nonproductive instead of productive complexes. Cellulose is also degraded by the synergistic action of cellulolytic enzymes, which also display this characteristic substrate inhibition (Va¨ ljama¨ e et al., 2001). Site-directed mutagenesis studies involving amino acids present in the putative catalytic site of M. sexta CHI have identified residues required for catalysis (Huang et al., 2000; Lu et al., 2002; Zhang et al., 2002). Aspartic acids 142 and 144, tryptophan 145, and glutamic acid 146 were identified as residues very important for catalysis and also for extending the pH range of enzyme activity into the alkaline pH range. Acidic and aromatic residues in other family 18 chitinases also are important for substrate binding and catalysis

129

(Watanabe et al., 1993, 1994; Uchiyama et al., 2001; Bortone et al., 2002). Some of these residues are essential only for crystalline chitin hydrolysis, whereas others are important not only for crystalline chitin hydrolysis but for other substrates as well (Watanabe et al., 2003). 4.3.4.7. Glycosylation of Insect Chitinases

Manduca sexta CHI is moderately N-glycosylated in the catalytic domain and heavily O-glycosylated in the linker region (Arakane et al., 2003). The insect cell line TN-5B1-4 (Hi 5), which is routinely used for expression of recombinant foreign glycoprotein, synthesizes proteins with both N- and O-linked oligosaccharides (Davidson et al., 1990; Davis and Wood, 1995; Jarvis and Finn, 1995; Hsu et al., 1997). Results of experiments investigating the effects of the N-glycosylation inhibitor tunicamycin on recombinant expression of insect chitinases in these cells indicated that the proteins were glycosylated prior to being secreted by the cells (Gopalakrishnan et al., 1995; Zheng et al., 2002). Direct chemical and enzymatic analyses confirmed that M. sexta CHI was both N- and O-glycosylated. Prolonged deglycosylation with a mixture of N- and O-glycosidases resulted in a protein that was smaller by about 6 kDa accounting for about 30 sugar residues per mole of protein (Arakane et al., 2003). Because N-linked oligosaccharides in insects typically have six or seven residues, two of which are GlcNAc (Paulson, 1989; Kubelka et al., 1995), the best estimate of the distribution of N-glycosylation indicated a single or possibly two sites of N-glycosylation in the catalytic domain and O-glycosylation of between 10 and 20 serine or threonine residues in the linker region. O-glycosylation may involve mainly addition of galactose and N-acetylgalactosamine. The chitinase from B. mori also is probably glycosylated because this protein and its breakdown product (65 kDa) stain with periodic acid–Schiff reagent. Further, the apparent mobility of the protein in sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) is 88 kDa, whereas the molecular weight of the mature protein predicted from the cDNA sequence is only 60 kDa (Koga et al., 1997). This protein has an S/T-rich linker similar to the M. sexta chitinase. On the other hand, the chitinase from wasp venom which has only a short linker region and is low in serine and threonine has nearly the same molecular weight as the one predicted from the cDNA sequence, suggesting that this protein may not be glycosylated (Krishnan et al., 1994). Thus, there is a good

130 Chitin Metabolism in Insects

correlation between the presence of an S/T-rich linker and extensive glycosylation (predominantly O-glycosylation) of the chitinolytic proteins. Glycosylation of the linker region may help to prevent proteolytic cleavage(s) at sites between the catalytic and chitin-binding domains. Such a functional role of glycosylated regions has been observed in some bacterial cellulases (Langsford et al., 1987). The full-length and near full-length O-glycosylated forms of M. sexta CHI were the most stable proteins when incubated with the midgut proteinases of the hornworm (Arakane et al., 2003). Protein modeling studies using the crystal structures of other family 18 glycosylhydrolases as templates suggested that the catalytic domain of M. sexta CHI has a (ba)8triose phosphate isomerase (TIM) barrel structure (Kramer and Muthukrishnan, 1997; Nagano et al., 2002). The ChBD probably exhibits a multistranded b-sheet structure based on similarity to tachycitin (Suetake et al., 2000). We know of no structures computed or proposed for linker domains, which may be very hydrophilic and rather flexible as well as potentially susceptible to proteolytic degradation unless they are protected by glycosylation. The CD spectrum of the linker domain was consistent with the lack of any secondary structure in this domain (Figure 5). It is conceivable that during the developmental period of maximum chitinase activity, the enzyme is fully glycosylated. When required, a glycosidase(s) could be produced that would remove sugar residues, thus exposing several more peptide bonds for proteolytic cleavage. Alternatively, proteolytic cleavage may be reduced because of glycosylation. Consistent with this notion is the finding that analysis of molting fluid from M. sexta and B. mori revealed the presence of truncated forms of catalytically active chitinases with sizes ranging from 50 to 60 kDa (Kramer and Koga, 1986; Koga et al., 1997; Abdel-Banat et al., 1999). We also detected similar truncated forms in our insect cell recombinant chitinase expression system, especially several days subsequent to infection with the recombinant baculovirus (Gopalakrishnan et al., 1995). 4.3.4.8. Antigenicity of Insect Chitinases

Invertebrate chitinases have been reported to elicit allergies in mammals. For example, a high prevalence of IgE antibodies to a tick chitinase was identified in canine atopic dermatitis with the chitinase formally designated Der f 15 (McCall et al., 2001). In ticks, this chitinase was localized in the proventriculus and intestine, indicating that it has a digestive, rather than molting-related, function. Like

insect chitinase, tick chitinase is extensively O-glycosylated on multiple sites along the 84 amino acid long S/T-rich sequence in the molecule. The transmission blocking antibody MF1 from the blood of gerbils infected with the nematode B. malayi was found to be directed against a microfilarial chitinase (Fuhrman et al., 1992). This antibody mediates the clearance of peripheral microfileremia in gerbils, indicating that chitinase is indeed a potent antigen. Even though it is unclear which region of the nematode chitinase is highly antigenic, the most probable one is the S/T–rich region known to be O-glycosylated. The primary epitope recognized by antibodies elicited by Manduca chitinases is the highly glycosylated S/T-rich linker region (Arakane et al., 2003). Other highly immunogenic insect proteins that also are extensively O-glycosylated in S/T-rich domains similar to the linker region of Manduca CHI are peritrophins-55 and -95 from the sheep blowfly, L. cuprina (Tellam et al., 2000, 2003). The sera of sheep vaccinated with these peritrophins exhibited a strong immune response that also inhibited growth of blowfly larvae (Casu et al., 1997; Tellam et al., 2003). 4.3.4.9. Other Possible Enzymes of Chitin Metabolism

Chitin deacetylases and chitosanases are two other enzymes that play major roles in chitin catabolism in other types of organisms. Chitin deacetylase catalyzes the removal of acetyl groups from chitin. This enzyme is widely distributed in microorganisms and may have a role in cell wall biosynthesis and in counteracting plant defenses (Tsigos et al., 2000). There is one report of an insect chitin deacetylase in physogastric queens of the termite Macrotermetes estherae (Sundara Rajulu et al., 1982). However, there have been no follow-up studies about this enzyme in other insect species. To our knowledge, there are no reports of chitosanases present in insects.

4.3.5. Nonenzymatic Proteins That Bind to Chitin There are approximately 32 families of CBDs that are defined as contiguous amino acid sequences within a carbohydrate-active enzyme or noncatalytic analogs, which exhibit a discrete fold having carbohydrate-binding activity (CAZY, 2004). One or more members in families 1, 2, 3, 5, 12, 14, 16, 18, and 19 are reported to bind chitin. Most, if not all, of the insect ChBDs, however, belong only to family 14.

Chitin Metabolism in Insects

Several chitinase-related proteins have been identified in insects, which are catalytically inactive because they are missing an amino acid residue critical for hydrolytic activity but nonetheless are carbohydrate-binding proteins with either a single copy or multiple repeats of ChBDs. These proteins may act as growth factors or play a defensive function as anti-inflammatory proteins. A chitinase homolog glycoprotein HAIP (hemolymph aggregation inhibiting protein) occurs in hemolymph of the lepidopteran M. sexta, which inhibits hemocyte aggregation (Kanost et al., 1994). A similar immunoreactive protein was detected in hemolymph of three other lepidopterans, B. mori, Heliothis zea, and Galleria mellonella. These proteins may have a role in modulating adhesion of hemocytes during defensive responses. Another glycoprotein, Ds47, which is produced in vitro by a Drosophila embryo-derived cell line and by fat body and hemocytes, may play a role in promoting the growth of imaginal discs (Kirkpatrick et al., 1995; Bryant, 2001). Another chitinase-related protein is induced together with a chitinase and b-N-acetylglucosaminidase by ecdysteroid in the anterior silk gland of B. mori at molting and at metamorphosis (Takahashi et al., 2002). The former is rather large in size and has a novel structure consisting of tandemly repeated catalytic domain-like plus linker sequences, but it has only one ChBD located in the middle of the protein. All of these proteins are evolutionarily related to chitinases, but they apparently have acquired a new growth-promoting or infection-resistance function that does not require catalytic activity. Evidently, chitinases have evolved into these lectin-like proteins by mutation of key residues in the active site, which abolishes enzyme activity and fine tunes the ligand-binding specificity. Chitin-binding proteins in vertebrates, invertebrates, and plants share a common structural motif composed of one to eight disulfide bonds and several aromatic residues, apparently the result of convergent evolution (Shen and Jacobs-Lorena, 1999; Suetake et al., 2000). A chitin-binding antifungal peptide from the coconut rhinoceros beetle, Oryctes rhinoceros, scarabaecin, is only 36 residues in length and contains only one disulfide bond (Hemmi et al., 2003). It shares significant tertiary structural similarity with ChBDs of other invertebrates and plants that have multiple disulfide bonds, even though there is no overall sequence similarity. Other invertebrate proteins that contain one or more ChBDs include the peritrophins (Tellam et al., 1999), mucins (Casu et al., 1997; Wang and Granados, 1997; Tellam et al., 1999; Rayms-Keller et al.,

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2000; Sarauer et al., 2003), and tachycitin (Suetake et al., 2000). Other proteins that bind to chitin include several lectins and cuticular proteins (see Chapter 4.2). The lectins are related to ChBDs found in PM and chitinases. Many insect cuticular proteins contain an amino acid sequence motif of approximately 35 residues known as the R&R consensus sequence (Rebers and Willis, 2001). This sequence, however, has no similarity to the cysteine-rich ChBDs found in chitinases, some PM proteins, and lectins. There are no or very few cysteine residues in the cuticular protein ChBDs (noncysChBD). Thus, there are two distinct classes of invertebrate ChBDs, those with the chitin-binding domain found in lectins, chitinases, and PM proteins (cysChBDs) and those with the cuticular protein chitin-binding domain (noncysChBDs). Homology modeling of insect cuticle proteins using the bovine plasma retinol binding protein as a template predicted an antiparallel bsheet half-barrel structure as the basic folding motif where an almost flat surface consisting of aromatic amino acid side chains interacts with the polysaccharide chains of chitin (Hamodrakas et al., 2002). In mammals there are several nonenzymatic members of the chitinase protein family. Oviduct-specific glycoprotein (OGP), a member of this family, is believed to be involved in the process of fertilization such as sperm function and gamete interactions (Araki et al., 2003). However, OGP was not essential for in vitro fertilization in mice, and so the functionality of OGP remains unknown. The human cartilage protein HCgp-39 is a chitin-specific lectin (Renkema et al., 1998; Houston et al., 2003) that is overexpressed in articular chondrocytes and certain cancers. It is thought to be an anti-inflammatoryresponse protein and/or to play a role in connective tissue remodeling. In contrast to chitinases, which bind and hydrolyze chitin oligosaccharides but do not undergo large conformational changes, HCgp39 exhibits a large conformational change upon ligand binding, which appears to signal the presence of chitinous pathogens such as fungi and nematodes (van Aalten, 2003). The murine Ym1 gene belongs to a family of mammalian genes encoding nonenzymatic proteins that are homologous to the chitinases from lower organisms, such as insects, nematodes, bacteria, and plants (Sun et al., 2001). YKL-40 is a nonenzymatic member of the mammalian family 18 glycosylhydrolases, which is a growth factor for connective tissue cells and stimulates migration of endothelial cells (Johansen et al., 2003). It is secreted in large amounts by

132 Chitin Metabolism in Insects

human osteosarcoma cells and murine mammary tumors, and it is also elevated in patients with metastatic breast cancer and colorectal cancer. These homologous mammalian proteins have no demonstrable chitinase activity and, therefore, cannot be considered chitinases. The biological functions of these proteins remain obscure. However, these proteins likely function through binding to carbohydrate polymers and since they are secreted from activated hemocytes, they may have a function in immunity such as a hemocyte inhibition (Falcone et al., 2001). Sequence comparison of these nonenzymatic and enzymatic proteins indicates that the enzymatic proteins have evolved into these lectins by the mutation of key residues in the active site and optimization of the substrate-binding specificity (Fusetti et al., 2002).

4.3.6. Regulation of Chitin Degradation The M. sexta chitinase and N-acetylglucosaminidase genes were shown to be upregulated by ecdysteroid (see Chapter 3.5) and down-regulated by the juvenile hormone mimic (see Chapter 3.7), phenoxycarb, in larval abdomens cut off from their hormonal sources (Fukamizo and Kramer, 1987; Koga et al., 1991; Kramer et al., 1993; Zen et al., 1996). Differential display was used to show that chitinase expression was regulated not only by ecdysteroid but also by juvenile hormone in the beetle T. molitor (Royer et al., 2002). Northern blot analysis of RNA from epidermis and 20-hydroxyecdysone-injected pupae showed that chitinase transcripts were correlated with molting hormone levels during metamorphosis. In addition, topical application of a juvenile hormone (JH) analog indirectly induced expression of chitinase mRNA. Thus, the Tenebrio chitinase gene is an early direct ecdysteroid-responsive one at the transcriptional level, but unlike M. sexta chitinase, it is apparently a direct target of JH as well. In the former case, the level at which JH regulates chitinase mRNA levels remains to be determined. The 20-hydroxyecdysone agonist, tebufenozide, induced expression of C. fumiferana chitinase when it was injected into mature larvae. The enzyme was produced 24 h post treatment in both the epidermis and molting fluid (Zheng et al., 2003).

4.3.7. Chitin Metabolism and Insect Control Chitinases have been used in a variety of ways for insect control and other purposes (Kramer et al., 1997; Gooday, 1999). Several chitinase inhibitors

with biological activity have been identified based on natural products chemistry (Spindler and Spindler-Barth, 1999), such as allosamidin (Rao et al., 2003) which mimics the carbohydrate substrate, and cyclic peptides (Houston et al., 2002). Although useful for biochemical studies, none of these chitin catabolism inhibitors have been developed for commercial use primarily because of their high cost of production and potential side effects. As we learn more details about chitinase catalysis, it might become more economically feasible to develop and optimize chitinase inhibitors for insect pest management. Additional uncharacterized steps in chitin synthesis and/or assembly of chitin microfibrils, on the other hand, have proved to be important for developing control chemicals that act selectively on economically important groups of insect pests (Verloop and Ferrell, 1977; Ishaaya, 2001). The benzoylphenylureas have been developed as commercial compounds for controlling agricultural pests. These antimolting insecticides are relatively nontoxic to mammals due to their strong protein binding and extensive metabolization to less toxic compounds (Bayoumi et al., 2003). Studies using imaginal discs and cell-free systems indicated that benzoylphenylureas inhibit ecdysteroid-dependent GlcNAc incorporation into chitin (Mikolajczyk et al., 1994; Oberlander and Silhacek, 1998). Those results suggest that benzoylphenylureas affect ecdysone-dependent sites, which leads to chitin inhibition. However, the site of action of the benzoylphenylureas still is not well known. Recently, several heteryl nucleoside nonhydrolyzable transition state analogs of UDP-GlcNAc were synthesized and evaluated for fungicidal activity, but they were not assayed for insecticidal activity (Behr et al., 2003). Entomopathogens secrete a plethora of extracellular proteins with potential activity in insect hosts. One of these proteins is chitinase, which is used by fungi such as Metarhizium anisopliae to help penetrate the host cuticle and render host tissues suitable for consumption (St. Leger et al., 1996; Krieger de Moraes et al., 2003). Among the 10 most frequent transcripts in a strain of M. anisopliae are three encoding chitinases and one a chitosanase, presumably reflecting a greater propensity to produce chitinases for host cuticle penetration (Freimoser et al., 2003a). Expressed sequence tag analysis of M. anisopliae may hasten gene discovery to enhance development of improved mycoinsecticides. However, when M. anisopliae was transformed to overexpress its native chitinase, the pathogenicity to the tobacco hornworm was unaltered, suggesting that

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wild-type levels of chitinase are not limiting for cuticle penetration (Screen et al., 2001). Another fungal species, Conidiobolus coronatus, also produces both endo- and exo-acting chitinolytic enzymes during growth on insect cuticle (Freimoser et al., 2003b). Apparently, both M. anisopliae and C. coronatus produce a chitinolytic enzyme system to degrade cuticular components. Both microbial and insect chitinases have been shown to enhance the toxicity of the entomopathogenic bacterium Bacillus thuringiensis (Bt) (Regev et al., 1996; Tantimavanich et al., 1997; Ding et al., 1998; Sampson and Gooday, 1998; Wiwat et al., 2000). For example, when the chitinolytic activities of several strains of B. thuringiensis were compared with their insecticidal activity, it was determined that the enzyme could enhance the toxicity of Bt to Spodoptera exigua larvae by more than twofold (Liu et al., 2002). Microbial chitinases have been used in mixing experiments to increase the potency of entomopathogenic microorganisms (review: Kramer et al., 1997). Synergistic effects between chitinolytic enzymes and microbial insecticides have been reported as early as the 1970s. Bacterial chitinolytic enzymes were first used to enhance the activity of Bt and a baculovirus. Larvae of C. fumiferana died more rapidly when exposed to chitinase–Bt mixtures than when exposed to the enzyme or bacterium alone (Smirnoff and Valero, 1972; Morris, 1976; Lysenko, 1976). Mortality of gypsy moth, Lymantria dispar, larvae was enhanced when chitinase was mixed with Bt relative to a treatment with Bt alone in laboratory experiments (Dubois, 1977). The toxic effect was correlated positively with enzyme levels (Gunner et al., 1985). The larvicidal activity of a nuclear polyhedrosis virus toward L. dispar larvae was increased about fivefold when it was administered with a bacterial chitinase (Shapiro et al., 1987). Chitin synthesis-inhibiting antifungal agents such as flufenoxuron and nikkomycin were used to promote the infection of silkworms with B. mori nucleopolyhedrovirus (Arakawa, 2002, 2003; Arakawa and Sugiyama, 2002; Arakawa et al., 2002). The mechanism of viral infection enhancement by these agents is not established, but it may involve destruction of PM structure, which would facilitate tissue invasion. Inducible chitinolytic enzymes from bacteria cause insect mortality under certain conditions. These enzymes may compromise the structural integrity of the PM barrier and improve the effectiveness of Bt toxin by enhancing contact of the toxin molecules with their epithelial membrane receptors. For example, five chitinolytic bacterial strains isolated from midguts of Spodoptera littoralis

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induced a synergistic increase in larval mortality when combined with Bt spore-crystal suspensions relative to either an individual bacterial strain or a Bt suspension alone (Sneh et al., 1983). An enhanced toxic effect toward S. littoralis also resulted when a combination of low levels of a truncated recombinant Bt toxin and a bacterial endochitinase was incorporated into a semisynthetic insect diet (Regev et al., 1996). Crude chitinase preparations from B. circulans enhanced the toxicity of Bt kurstaki toward diamondback moth larvae (Wiwat et al., 1996). Liu et al. (2002) recently reported that several strains of Bt produced their own chitinases, which had synergistic larvicidal activity with the endotoxins. In biopesticide development research, we used a family 18 insect chitinase as an enhancer protein for baculovirus toxicity and as a host plant resistance factor in transgenic plants. Introduction of an insect chitinase cDNA into A. californica multiple nuclear polyhedrosis viral (AcMNPV) DNA accelerated the rate of killing of fall armyworm compared to the wild-type virus (Gopalakrishnan et al., 1995). Baculoviral chitinases themselves play a role in liquefaction of insect hosts (Hawtin et al., 1997; Thomas et al., 2000). A constitutively expressed exochitinase from B. thuringiensis potentiated the insecticidal effect of the vegetative insecticidal protein Vip when they were fed to neonate larvae of S. litura (Arora et al., 2003). Some granuloviruses, on the other hand, do not utilize chitinases in a similar manner, which helps to explain why some granulovirus-infected insects do not lyse at the end of the infection process (Wormleaton et al., 2003). Mutagenesis of the AcMNPV chitinase gene resulted in cessation of liquefaction of infected T. ni larvae, supporting a role of chitinase in virus spread (Thomas et al., 2000). However, the insecticidal activity of insect chitinase was not substantial enough for commercial development. We have attempted with little success to improve the catalytic efficiency and stability of this enzyme so that its pesticidal activity would be enhanced (Lu et al., 2002; Zhang et al., 2002; Arakane et al., 2003). Nevertheless, tobacco budworms were killed when reared on transgenic tobacco expressing a truncated, enzymatically active form of insect chitinase (Ding et al., 1998). We also discovered a synergistic interaction between insect chitinase expressed in transgenic tobacco plants and Bt (applied as a spray at sublethal levels) using the tobacco hornworm as the test insect. In contrast to results obtained with the tobacco budworm, studies with the hornworm revealed no consistent differences in larval growth or foliar damage when the insects were reared on

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first-generation transgenic chitinase-positive tobacco plants as compared to chitinase-negative control plants. When Bt toxin was applied at levels where no growth inhibition was observed on control plants, chitinase-positive plants had significantly less foliar damage and lower larval biomass production. These results indicated that the insect chitinase transgene did potentiate the effect of sublethal doses of Bt toxin and vice versa (Ding et al., 1998). Tomato plants have been transformed with fungal chitinase genes with concomitant enhancement in resistance to insect pests (Gongora et al., 2001). Effects observed include reduced growth rates and increased mortality, as well as a decrease in plant height and flowering time with an increase in the number of flowers and fruits (Gongora and Broadway, 2002). Chitinase-secreting bacteria have been used to suppress herbivorous insect pests. A chitinase gene-transformed strain of Enterobacter cloacae digested the chitinous membranes of phytophagous ladybird beetles, Epilachna vigintioctopunctata, and also suppressed leaf-feeding and oviposition when the beetles ingested transformed bacteria entrapped in alginate microbeads sprayed on tomato seedlings (Otsu et al., 2003). Several GlcNAc-specific lectins from plants have been evaluated for insect toxicity (Harper et al., 1998; Macedo et al., 2003). These proteins appear to disrupt the integrity of the PM by binding to chitin or glycan receptors on the surface of cells lining the insect gut. They also may bind to glycosylated digestive enzymes and inhibit their activity. Another type of plant chitin-binding protein is the seed storage protein, vicilin, which is actually a family of oligomeric proteins with variable degrees of glycosylation (Macedo et al., 1993; Shutov et al., 1995). Some vicilins are insecticidal to bruchid beetles and stalk borers (Sales et al., 2001; Mota et al., 2003). Apparently, these proteins bind to the PM, causing developmental abnormalities and reduced survival rates. To date no carbohydrate-binding protein derived from an insect has been evaluated for biocidal activity. A novel approach has been proposed to develop strategies for insect control by utilizing chitin-binding molecules to specifically target formation of the PM. Calcofluor, a chemical whitener with chitin-binding properties, was used as a model compound in the diet to inhibit PM formation in T. ni and also to increase larval susceptibility to baculovirus infection (Wang and Granados, 2000b). It also was effective in suppressing PM formation in Spodoptera frugiperda and at the same time in preventing the establishment of a decreasing gradient of proteinases along the midgut tissue (Bolognesi et al., 2001).

Another type of hydrolytic enzyme with a ChBD has been shown to exhibit insecticidal activity in plants. Maize accumulates a 33 kDa cysteine protease containing a ChBD in response to insect feeding (Perchan et al., 2002). This enzyme apparently damages the insect’s PM by utilizing the ChBD to localize itself at the chitin-protein-rich PM, where the PM proteins are digested, rendering the PM dysfunctional. Another protease with a chitin-binding domain has been described from A. gambiae, which may be involved in insect defense (Danielli et al., 2000). This 147 kDa protein, sp22D, is expressed in a variety of tissues, most strongly in hemocytes, and is secreted into the hemolymph. Upon bacterial infection, the transcripts for this protein increase by about twofold suggesting a role in insect defense. This protein has a multidomain organization that includes two copies of an N-terminal ChBD, a C-terminal protease domain, and additional receptor domains. It binds strongly to chitin and undergoes complex proteolytic processing during pupal to adult metamorphosis. It has been proposed that exposure of this protease to chitin may regulate its activity during tissue remodeling or wounding. Recently, two synthetic peptides were found to inhibit A. gambiae midgut chitinase and also to block sporogonic development of the human malaria parasite, Plasmodium falciparum, and avian malaria parasite, P. gallinaceum, when the peptides were fed to infected mosquitoes (Bhatnagar et al., 2003). The design of these peptides was based on the putative proregion sequence of mosquito midgut chitinase. The results indicated that expression of chitinase inhibitory peptides in transgenic mosquitoes might alter the vectorial capacity of mosquitoes to transmit malaria.

4.3.8. Concluding Remarks Although chitin was discovered nearly two centuries ago, it remains a biomaterial in waiting because, unlike other natural materials such as collagen and hyaluronic acid, very few technological uses have been developed (Khor, 2002; Tharanathan and Kittur, 2003). There are many unanswered questions about chitin morphology and chitin deposition in the insect cuticle and PM. We do not know how or whether chitin forms covalent interactions with other components in these extracellular matrices. Chitosan, on the other hand, does react with quinones (Muzzarelli and Muzzarelli, 2002; Muzzarelli et al., 2003). Thus, if there were any free amino groups in insect chitin, C–N linkages between chitin and catechols would be expected (Schaefer et al.,

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1987). We do not yet understand how factors such as metal ions affect chitin metabolism. In fungi, ions such as zinc were found to alter chitin deposition and morphology (Lanfranco et al., 2002). Perhaps, in insects there is an ionic effect on differential expression of CHS isozymes. We know much more about insect chitinolytic enzymes than about insect chitin biosynthetic enzymes. Many questions remain about the biosynthesis of insect chitin, not the least of which are why insects have multiple genes for CHS, how many CHSs are required to make an insect, at what developmental stages are the various CHSs produced, and what are the unique properties and functions of each CHS. Of particular interest is the role of alternate splicing in generating different isoforms of CHSs from the same gene. The developmental cues that control alternate splicing and how they affect chitin synthesis and/or deposition will be subjects of future studies. The cloning of CHS genes should soon lead to availability of large amounts of recombinant enzymes or subdomains thereof using appropriate expression systems. Studies with pure proteins and the availability of molecular probes will provide a better understanding of the chitin biosynthetic pathway and its regulation in the future. Two other major questions about insect chitin biosynthesis are: what is the mechanism of the initiation phase and is there an autocatalytic initiator. Like glycogen synthesis, chitin synthesis probably includes both initiation and elongation phases. As the initiator of glycogen synthesis, glycogenin transfers glucose from UDP-glucose to itself to form an oligosaccharide–protein primer for elongation (Gibbons et al., 2002). Like chitin synthase, glycogenin is a glycosyltransferase, which raises the question of whether chitin synthase has an autocatalytic function similar to glycogenin and whether there is a chitinogenin-like protein. Another possibility is the participation of a lipid primer for chitin synthesis. Recently, cellulose synthesis in plants was found to involve the transfer of lipidlinked cellodextrins to a growing glucan chain (Read and Bacic, 2002). The lipid in this case was sitosterol-b-glucoside. Little is known about the catalytic mechanism of any insect CHS. Once insect CHS-related recombinant proteins are obtained, site-directed mutagenesis can be used to probe for essential residues in the catalytic and regulatory domains. It is likely that acidic amino acids play critical roles in CHS catalysis in a manner comparable to those identified in other glycosyltransferases (Hefner and Stockigt, 2003) and in yeast chitin synthases (Nagahashi et al., 1995).

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Chitinolytic enzymes are gaining importance for their biotechnological applications in agriculture and healthcare (Patil et al., 2000). Additional success in using chitinases for different applications depends on a better understanding of their biochemistry and regulation so that their useful properties can be optimized through genetic and biochemical engineering. Reasons for the rather high multiplicity of domain structures for insect and other chitinases are not fully understood. So far little success has occurred in using chitinase in pest control applications, but it may prove more useful as an enhancer protein in a cocktail with other biopesticides targeted at the cuticle or gut. Also, only a few catalytic domains or chitin-binding domains or various combinations thereof have been evaluated for biocidal activity and thus, further toxicological experimentation is warranted. Although substantial progress in studies of insect chitin metabolism has occurred since the first edition of Comprehensive Insect Physiology, Biochemistry, and Pharmacology was published in 1985, we still do not know much about how chitin is produced and transported across the membrane so that it can interact perfectly with other components for assembly of the supramolecular extracellular structures called the exoskeleton and PM. These materials are still very much biochemical puzzles in which we do not understand well how the various components come together during morphogenesis or are digested apart during the molting process. Hopefully, this chapter will stimulate more effort to understand how insects utilize chitin metabolism for growth and development, and to develop materials that may perturb insect chitin metabolism for pest management purposes.

Acknowledgments The authors are grateful to Sherry Mowbray, Wimal Ubhayasekera, Yasuyuki Arakane, Qingsong Zhu, David Hogenkamp, Renata Bolognesi, Tamo Fukamizo, Daizo Koga, Walter Terra and Clelia Terra for their help with the preparation and/or comments on various aspects of this review. Supported in part by National Science Foundation grant no. 0316963. Mention of a proprietary product does not constitute a recommendation or endorsement by the US Department of Agriculture. The US Department of Agriculture is an equal opportunity/affirmative action employer and all agency services are available without discrimination. This is contribution no. 04-058-C of the Kansas Agricultural Experiment Station.

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4.4 Cuticular Sclerotization and Tanning S O Andersen, University of Copenhagen, Copenhagen, Denmark ß 2005, Elsevier BV. All rights reserved.

4.4.1. Introduction 4.4.2. Current Model for Cuticular Sclerotization 4.4.3. Sclerotization (Tanning) Precursors 4.4.3.1. N-Acetyldopamine and N-b-Alanyldopamine 4.4.3.2. Putative Sclerotization Precursors 4.4.3.3. Noncuticular Sclerotization 4.4.4. Cuticular Enzymes and Sclerotization 4.4.4.1. ortho-Diphenoloxidases 4.4.4.2. Laccases 4.4.4.3. Cuticular Peroxidases 4.4.4.4. ortho-Quinones and para-Quinone Methides 4.4.4.5. Dehydro-NADA and Dehydro-NBAD 4.4.4.6. Various Catechol Derivatives Obtained from Cuticles 4.4.5. Control of Sclerotization 4.4.5.1. Pre-Ecdysial Sclerotization 4.4.5.2. Post-Ecdysial Sclerotization 4.4.5.3. Puparial Sclerotization 4.4.5.4. Transport of Sclerotization Precursors to the Cuticle 4.4.5.5. Balance between Cuticular Enzymes 4.4.5.6. Intensity of Cuticular Sclerotization 4.4.6. Comparative Aspects 4.4.6.1. Cuticular Darkening 4.4.6.2. Cuticular Sclerotization in Insects Compared to That in Other Arthropods 4.4.7. Unsolved Problems 4.4.7.1. Alternative Pathway for Dehydro-NADA Formation? 4.4.7.2. Extracuticular Synthesis of Catechol–Protein Conjugates for Sclerotization? 4.4.7.3. Importance of Cuticular Dehydration? 4.4.7.4. Lipids and Sclerotization?

4.4.1. Introduction The cuticle covers the complete body of the insects as an effective barrier between the animal and its surroundings; it provides protection against desiccation, microorganisms, and predators, and as an exoskeleton it serves as attachment sites for muscles. Cuticle is typically divided into relatively hard and stiff regions, the sclerites, separated by more flexible and pliable regions, the arthrodial membranes, which make the various forms of locomotion possible. Marked differences in mechanical properties may also be present on the microscopical level; two neighboring epidermal cells can produce cuticle with highly contrasting properties, indicating a precise control of cuticular composition. The mechanical properties of individual cuticular regions correspond closely to their functions and the forces to which they are exposed during the normal life of the animal. Proper flight can thus only be sustained when all wing regions have near-optimal balance

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between stiffness and flexibility; if the wing material locally is too soft or too stiff, the varying air pressure during the wing strokes will not cause the wings to bend to the shapes needed for generating optimal lift. The mechanical properties of cuticle are determined by the interplay of many factors, such as cuticular thickness, relative amounts of chitin and proteins, chitin architecture, protein composition, water content, intracuticular pH, and degree of sclerotization and other secondary modifications. Sclerotization of insect cuticle has been reviewed several times in recent years (Sugumaran, 1988, 1998; Andersen, 1990; Hopkins and Kramer, 1992; Andersen et al., 1996), but many aspects of the process are still rather poorly understood. Besides giving an overview of the present knowledge of the sclerotization process attention is drawn to some problems that need to be investigated in more detail.

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Cuticular sclerotization is a chemical process whereby certain regions of insect cuticle are transformed irreversibly from a pliant material into a stiffer and harder structure, characterized by decreased deformability, decreased extractability of the matrix proteins, and increased resistance towards enzymatic degradation. During sclerotization the color of the cuticle may change from nearly colorless over various brown and black shades to the completely black. The term tanning is often used synonymously with sclerotization, but sometimes it is specifically used for the processes resulting in light brown (tan) cuticles. Sclerotization often takes place in connection with molting, starting just after the new, yet unsclerotized cuticle has been expanded to its final size and shape, but some specialized cuticular regions are sclerotized while the insect is still in its pharate state inside the old cuticle. Such pre-ecdysially sclerotized regions cannot be expanded post-ecdysially, but may help the insect to escape from the exuvium. The dipteran puparium is an example of a soft larval cuticle, which is sclerotized at the end of the last larval instar to form a hard protective case inside which metamorphosis to pupa and adult can take place. Sclerotization of structural materials in insects is not restricted to cuticle; other materials as well, such as egg cases and silks, may be stabilized by chemical processes closely related to cuticular sclerotization.

4.4.2. Current Model for Cuticular Sclerotization During the years several models have been proposed for the chemical reactions which occur in the insect cuticle during the sclerotization process, and although many details of the individual steps in the reactions are still controversial or unexplored, there is general agreement concerning the main features of the process. The currently accepted sclerotization model is shown in Figures 1 and 2, and its main features are: the amino acid tyrosine (1) is hydroxylated to 3,4-dihydroxyphenylalanine (DOPA, 2), which by decarboxylation is transformed to dopamine (3), a compound of central importance for both sclerotization and melanine formation. Dopamine can be N-acylated to either N-acetyldopamine (NADA, 4) or N-b-alanyldopamine (NBAD, 5), and both can serve as precursors in the sclerotization process. They are enzymatically oxidized to the corresponding o-quinones (6), which can react with available nucleophilic groups, whereby the catecholic structure is regained and the nucleophile is linked to the aromatic ring (11). The o-quinones of NADA and NBAD may also be enzymatically isomerized to the corresponding p-quinone methides (7), and the b-position of their side chain react readily with nucleophiles (12). The p-quinone methides may be enzymatically isomerized to side chain unsaturated catechol derivatives (8), dehydro-NADA

Figure 1 Biosynthesis of the sclerotization precursors NADA and NBAD from tyrosine. The enzyme tyrosine hydroxylase hydroxylates tyrosine (1) to 3,4-dihydroxyphenylalanine (DOPA, 2), which is decarboxylated to dopamine (3) by the enzyme dopa-decarboxylase. Dopamine can be enzymatically acylated to either N-acetyldopamine (NADA, 4) or N-b-alanyldopamine (NBDA, 5). Surplus dopamine can be used for melanin synthesis.

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147

Figure 2 Cuticular oxidation of NADA and NBAD to various sclerotization agents. The acyldopamines are enzymatically oxidized to o-quinones (6), which can react spontaneously with nucleophilic compounds (HX) to give ring-substituted adducts (11), or they can be isomerized to the more reactive p-quinone methides (7), which can react with nucleophiles to give side chain substituted adducts (12). The quinone methides may also be isomerized to dehydro-acyldopamines (8), which after oxidation to quinones (9 and 10) can react with catechols to give dihydroxyphenyl-dihydrobenzodioxine derivatives (13).

and dehydro-NBAD, which after oxidation to unsaturated quinones can react with catechols to form dihydroxyphenyl-dihydrobenzodioxine derivatives (13) and with other nucleophilic groups to give yet unidentified compounds. The o-quinones and p-quinone methides react preferably with the imidazole group in the cuticular proteins, but may also react with free amino groups, such as terminal amino groups in proteins and e-amino groups in lysine residues, with catechols, with water, and probably also with hydroxyl groups in the N-acetylglucosamine residues in chitin. Depending upon the degree of sclerotization the various reactions between quinones and nucleophilic residues in the cuticular matrix proteins will result in the proteins being more or less covered by aromatic residues; some of these residues may be involved in cross-linking the cuticular proteins and maybe also forming links between proteins and chitin; some will be linked to only a single protein molecule, thereby increasing its hydrophobicity without being part of a covalent cross-link. During sclerotization most of the water-filled spaces between the matrix

proteins in the presclerotized cuticle will become filled with polymerized catecholic material. As a result of these processes the interactions between the cuticular components have become stronger, the peptide chains more difficult to deform, and the proteins cannot be moved relative to each other or to the chitin filament system. Together, all these changes contribute to make the material stiffer and more resistant towards degradation. The various reactions involved in the model will be discussed in more detail in the following sections, with main emphasis on aspects where the evidence is insufficient or missing or where some observations disagree with the scheme, to indicate areas where more research is needed. The appearance and properties of cuticle from different body regions of the same animal can vary widely, and a considerable part of this variation, such as the different coloration and mechanical properties, is probably due to quantitative or qualitative differences in the sclerotization process. There is no compelling reasons to believe that exactly the same stabilization process is used for sclerotization in all types of solid cuticle,

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and generalizations based upon results obtained with a single or a few insect species can easily be misleading. A number of cuticular types will have to be analyzed to determine how the individual steps involved in sclerotization are modulated to give the local variation between cuticular regions of a given insect and between cuticles from different insect species. It will also be important to study how the reactions are controlled to give an optimal degree of sclerotization. Most of the results and ideas presented in this chapter have been obtained by studies involving material from only a few insect species, such as cuticle of blowfly larvae, pupae of Manduca sexta, and locust femurs, and it is to be expected that detailed studies of cuticle from more species will give a much more varied and fascinating picture of the complexity of cuticular sclerotization.

4.4.3. Sclerotization (Tanning) Precursors The terms sclerotization agents and tanning agents were originally used for the compounds which are secreted from the epidermal cells into the cuticle, where they are oxidized by enzymes to become sufficiently reactive to form covalent linkages to proteins and chitin. There is now a tendency to restrict the term sclerotization agents to the reactive species directly involved in forming links to the cuticular components. The compounds secreted from the epidermis to be activated in the cuticle shall accordingly be called sclerotization or tanning precursors (Sugumaran, 1998). 4.4.3.1. N-Acetyldopamine and N-b-Alanyldopamine

The first discovered and most common precursor for cuticular sclerotization is NADA (4), which is synthesized by N-acetylation of dopamine. The central role of NADA in sclerotization was demonstrated by Karlson’s research group during studies of tyrosine metabolism in insects (review: Karlson and Sekeris, 1976). They showed that NADA is incorporated into the puparial cuticle of the blowfly Calliphora vicina during its sclerotization, and that radioactively labeled tyrosine was metabolized to NADA when injected into last instar larvae shortly before puparium formation, and degraded when injected into younger larvae. NADA was also shown to be involved in cuticular sclerotization in several other insect species, such as the desert locust, Schistocerca gregaria (Karlson and SchlossbergerRaecke, 1962; Schlossberger-Raecke and Karlson, 1964). Incorporation of NADA into cuticle can be a very efficient process; after injection of radioactive

NADA into young adult locusts about 80% of the total radioactivity was later recovered from the sclerotized cuticle (Andersen, 1971). NADA appears to be involved in cuticular sclerotization in all insect species investigated. The amino acid b-alanine has been reported as a constituent of several types of sclerotized cuticle, and was suspected to participate somehow in the sclerotization process (Andersen, 1979a). Hopkins et al. (1982) showed that the b-alanyl derivative of dopamine, NBAD (6), is a sclerotizing precursor in the cuticle of M. sexta pupae, thus accounting for the presence of b-alanine in hydrolysates of the fully sclerotized cuticle. NBAD is also a sclerotization precursor in other cuticles from which b-alanine is released by acid hydrolysis, such as the cuticle of the red flour beetle, Tribolium castaneum (Kramer et al., 1984). The synthesis and utilization of NBAD during pupation of M. sexta has been reported (Krueger et al., 1989). An enzyme system in the medfly, Ceratitis capitata which can catalyze b-alanylation of dopamine to give NBAD has been partially characterized (Pe´rez et al., 2002). The two sclerotizing compounds, NADA and NBAD, are used together in many types of cuticles; but the cuticle of some insects, such as the locusts, S. gregaria and Locusta migratoria, appears to be exclusively sclerotized by NADA, as no b-alanine has been obtained from their acid hydrolysates. No cuticles have yet been reported to be sclerotized exclusively by NBAD. A correlation appears to exist between the intensity of brown color of the fully sclerotized cuticle and the amounts of NBAD taking part in the sclerotization process: cuticles that are sclerotized exclusively by NADA are colorless or very lightly straw-colored, and the more NBAD dominates in the process the darker brown will the cuticle become (Brunet, 1980; Hopkins et al., 1984). Czapla et al. (1990) reported that cuticular strength in five differently colored strains of the cockroach Blattella germanica correlated well with their concentrations of b-alanine and NBANE, whereas dopamine concentration correlated with melanization. Cuticular strength as well as cuticular concentrations of b-alanine and NBAD increased more rapidly in the rust-red wild-type of T. castaneum than in the black mutant strain, whereas cuticular dopamine increased more rapidly in the black strain than in the wild-type (Roseland et al., 1987). Significant amounts of sclerotization precursors are often present as conjugates before the onset of sclerotization. The conjugates, which can be glucosides, phosphates, or sulfates (Brunet, 1980; Kramer and Hopkins, 1987), are not easily oxidized, and

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have to be hydrolyzed to free catechols before they can take part in sclerotization. It is assumed that the catechol conjugates serve as a storage reservoir of catecholamines ready to be used when the need for sclerotization arises (Brunet, 1980). A dopamine conjugate, identified as the 3-O-sulfate ester, is present in the hemolymph of newly ecdysed cockroaches, and its concentration decreases rapidly as the cockroach cuticle sclerotizes. The sulfate moiety is not transferred into the cuticle, and removal of sulfate and acylation of the liberated dopamine to NADA and/or NBAD most likely occurs in the epidermal cells (Bodnaryk and Brunet, 1974; Czapla et al., 1988, 1989). Hopkins et al. (1984) reported that a large fraction of the various catecholamines in M. sexta hemolymph and cuticle is present as acid labile conjugates. In larval and pupal hemolymph these conjugates are mainly 3-O-glucosides together with small amounts of the 4-O-glucosides, whereas adult hemolymph contains more of the 4-O-glucoside than of the 3-O-glucoside (Hopkins et al., 1995). Both conjugated and unconjugated forms of NADA and NBAD were extracted from M. sexta cuticle (Hopkins et al., 1984), indicating that the epidermal cells possess transporting systems for conjugated as well as unconjugated catecholamines. It is also likely that a b-glucosidase activity is present inside the Manduca cuticular matrix, to catalyze the release of the sclerotization precursors from their conjugates, as the conjugated forms cannot be used directly for sclerotization. 4.4.3.2. Putative Sclerotization Precursors

So far, convincing evidence that they function as cuticular sclerotization precursors has only been obtained for NADA and NBAD, but several other compounds have been described as likely sclerotization precursor candidates, such as dopamine (3), N-acetyl-norepinephrine (NANE) (14), N-balanyl-norepinephrine (NBANE) (15), and 3,4dihydroxyphenylethanol (DOPET) (16) (Figure 3).

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It is likely that they all have some role to play in sclerotization. 4.4.3.2.1. Dopamine Dopamine is precursor for both NADA and NBAD synthesis, and it can also be precursor for black, insoluble melanins. Melaninlike materials are of common occurrence in cuticular structures, and they can be present either in microscopic granules or homogeneously distributed within the cuticular matrix (Kayser-Wegmann, 1976; KayserWegmann and Kayser, 1983; Hiruma and Riddiford, 1988). The granules are produced within epidermal cells and transported to the subepicuticular space via long cellular projections (Curtis et al., 1984; Kayser, 1985), and apparently the melanin in the granules is linked to granular proteins, but not to proteins in the cuticular matrix. The diffusely distributed melanins appears to be formed in situ within the cuticular matrix, and covalent links are probably formed between the polymeric melanin and the matrix proteins, thereby rendering the proteins more stable and insoluble, contributing to both darkening and increased mechanical stiffness of the cuticle. It will therefore be difficult in all cases to discern between the process of melanization of cuticle and the process of sclerotization. The observation that cuticular incorporation of radioactive tyrosine is nearly the same in wild-type and albino mutant of S. gregaria (Karlson and Schlossberger-Raecke, 1962) shows that melanization does not play a major role in sclerotization, but handling of small samples of melanized or albino cuticle indicates that it may have a minor role, as the melanized samples appear to be more brittle than the unmelanized (S.O. Andersen, unpublished data). A quantitative comparison of the mechanical properties of the two cuticular samples, to see to what extent melanization influences the physical properties of the material, would be interesting. Melanin can be formed from either DOPA or dopamine by slightly different routes. DOPA can via dopachrome and 5,6-dihydroxyindole carboxylic acid be transformed to 5,6-dihydroxyindole,

Figure 3 Putative precursors for cuticular sclerotization. 14, N-acetylnorepinephrine (NANE); 15, N-b-alanylnorepinephrine (NBANE); 16, 3,4-dihydroxyphenylethanol (DOPET); 17, gallic acid.

150 Cuticular Sclerotization and Tanning

which after enzymatic oxidation polymerizes to melanin, and transformation of dopamine to 5,6dihydroxyindole goes via dopamine chrome. The formation of the two intermediates, dopachrome and dopamine chrome, is catalyzed by different, but related enzymes. Dopamine appears to be a better substrate than DOPA for formation of cuticular melanin, and the melanins formed in hemolymph during defense reactions appear to be formed mainly via the DOPA pathway. In Drosophila melanogaster the products of two of the yellow genes, yellow-f and yellow-f2, are dopachrome-conversion enzymes, which have low activities towards dopamine chrome. The product of the gene yellow-y, involved in wing and cuticle melanization, has significant sequence similarity to yellow-f and yellow-f2 proteins, but is apparently devoid of dopachromeconversion activity (Han et al., 2002). Whether the yellow-y protein can catalyze the conversion of dopamine chrome to 5,6-dihydroxyindole was not reported. The black body color of the D. melanogaster mutants black and ebony is due to the inability of these mutants to produce sufficient NBAD for cuticular sclerotization: black is defective in the synthesis of b-alanine, and can be rescued by injection of b-alanine, and ebony is defective in the enzyme NBAD-synthetase, and cannot be rescued by injection of b-alanine (Wright, 1987). The result is that in both mutants some of the dopamine not used for NBAD synthesis is channeled into cuticular melanin production. Both stiffness and puncture resistance of the cuticle are decreased in the mutants, and electron microscope studies show that the cuticular chitin lamellae are abnormally wide and diffuse (Jacobs, 1978, 1980, 1985), indicating that even if dopamine-derived melanin can take part in cuticular stabilization, the result is inferior to the material obtained by NBAD sclerotization. The tan mutant of D. melanogaster, which is characterized by absence of the wild-type cuticular melanin pattern, has low activity of the enzyme N-b-alanyldopamine-hydrolase, catalyzing the hydrolysis of NBAD to b-alanine and dopamine (Wright, 1987). The enzyme systems, responsible for NBAD synthesis and NBAD hydrolysis, respectively, are probably located in different compartments, and the presence of melanin in some but not all cuticular regions of wild-type fruitflies could be explained by the local presence of the NBAD-hydrolase within the matrix of the melanizing regions, where it will hydrolyze some of the NBAD secreted from the epidermal cells, creating a dopamine concentration sufficient to stimulate a localized melanin production.

4.4.3.2.2. N-acetylnorepinephrine (NANE) and N-b-alanylnorepinephrine (NBANE) NANE and NBANE are special cases among the cuticular catechols, as they can be considered both as byproducts of the sclerotization process and precursors for sclerotization. They have been reported to occur both free and as an O-glucoside in hemolymph and integument in several insects (Hopkins et al., 1984, 1995; Morgan et al., 1987; Czapla et al., 1989). NANE and NBANE can be generated within the cuticle, when the enzymatically produced p-quinone methides of NADA and NBAD react with water instead of reacting with cuticular proteins or isomerizing to dehydro-derivatives, but they may also be produced by hydrolysis of some unidentified products of the sclerotization process. Mild acid treatment of sclerotized cuticles can release some NANE and NBANE from the cuticular structure, probably due to hydrolysis of a bond between the b-position of the catechols and some cuticular constituent. The nature of the bond is uncertain, but it could well be an ether linkage connecting the acyldopamine side chain to chitin. Formation of an ether, b-methoxyNADA, occurs when isolated pieces of cuticle or extracted cuticular enzymes act upon NADA in the presence of methanol; the compound is acid labile and is readily hydrolyzed to free NANE (Andersen, 1989c; Sugumaran et al., 1989b). NANE can be covalently incorporated into the cuticular matrix during sclerotization, indicating that the compound can serve as a sclerotization precursor (Andersen, 1971), and this is probably also the case for NBANE. When radioactively labeled NANE was injected into newly ecdysed locusts a significant fraction of the radioactivity (about 10%) was incorporated into the cuticle, and hydrolysis of the cuticle was needed to release the activity. Acid hydrolysis of cuticle from locusts injected with labeled norepinephrine resulted in the release of both labeled norepinephrine and arterenone, whereas little radioactivity was present in the neutral ketocatechol fraction. This is in contrast to parallel experiments when labeled dopamine was injected and nearly all the radioactivity was recovered as neutral ketocatechols, indicating that the cuticular enzymes can catalyze the incorporation of norepinephrine and NANE into the cuticular matrix, but not as efficiently and not by the same route as the incorporation of dopamine and NADA. 4.4.3.2.3. Dihydroxyphenylethanol (DOPET) The third putative sclerotization precursor, 3,4-dihydroxyphenylethanol (DOPET), is also present in the hemolymph and integument of insects; it has been obtained from cuticle of the cockroach Periplaneta

Cuticular Sclerotization and Tanning

americana (Atkinson et al., 1973b; Czapla et al., 1988) and the beetle Pachynoda sinuata (Andersen and Roepstorff, 1978), and it can function as substrate for the cuticular phenoloxidases. Extraction and acid hydrolysis of sclerotized cuticles have yielded various DOPET derivatives, suggesting that DOPET is transported into the cuticle and incorporated into the cuticular matrix during sclerotization. Adducts of DOPET and histidine have been obtained by acid hydrolysis of sclerotized Manduca pupal cuticle and identified by means of mass spectrometry (Kerwin et al., 1999). A dihydroxyphenyl-dihydrobezodioxine-type adduct of DOPET and NADA has also been extracted from sclerotized beetle (P. sinuata) cuticle (Andersen and Roepstorff, 1981), but the metabolism of DOPET in insects has not been studied in much detail. The relative roles of dopamine, NANE, NBAD, and DOPET compared to the two major sclerotization precursors, NADA and NBAD, have never been properly established. The compounds are probably only of minor importance for the mechanical properties of sclerotized cuticles, but their involvement in the sclerotization process may play a role in fine-tuning of the cuticular properties. 4.4.3.2.4. Other sclerotization precursors It has been reported that improved growth of the tree locust, Anacridium melanorhodon can be obtained

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by addition of gallic acid (17) and other plant phenols to its food, and that the ingested plant phenols are incorporated into the cuticle and may contribute to its stabilization (Bernays et al., 1980; Bernays and Woodhead, 1982). 4.4.3.3. Noncuticular Sclerotization

Although the sclerotization process has mainly been studied in the cuticle of insects, related processes are used for stabilization of other insect materials, such as silks, egg capsules, and chorions, and their structures are shown in Figure 4. Pryor (1940a) showed that 3,4-hydroxybenzoic acid (18) is a sclerotization precursor for egg capsules of the cockroach Blatta orientalis, and Pau and Acheson (1968) reported that other cockroach species use 3,4-dihydroxybenzyl alcohol (19) as a precursor for egg capsule sclerotization. The praying mantid, Hierodula patellifera uses various catechol derivatives for stabilizing the egg capsules; besides NADA and NBAD the following catechols have been reported: N-malonyldopamine (20), N-(N-acetyl-b-alanyl) dopamine (21), and N-(N-malonyl-b-alanyl)dopamine (22) (Kawasaki and Yago, 1983; Yago and Kawasaki, 1984; Yago et al., 1984). The cockroach catechols are stored in the left colleterial gland as 4-O-glucosides (Brunet, 1980), and the catechols in praying mantids are stored as 3-O-glucosides (Yago

Figure 4 Precursors for noncuticular sclerotization. 18, 3,4-dihydroxybenzoic acid (protocatechuic acid); 19, 3,4-dihydroxybenzyl alcohol; 20, N-malonyldopamine; 21, N-(N-acetyl-b-alanyl)dopamine; 22, N-(N-malonyl-b-alanyl)dopamine; 23, gentisic acid; 24, 3-hydroxyanthranilic acid; 25, N-(3,4-dihydroxyphenyllactyl)DOPA.

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et al., 1984). The right colleterial gland in both species contains a glucosidase, and when the secretions from the two glands are mixed, the glucosides are hydrolyzed, and the liberated catechols are enzymatically oxidized to o-quinones, which in mantids may be further converted to a highly reactive intermediates, presumably p-quinone methides (Yago et al., 1990). The eggs of the grasshopper Melanoplus sanguinipes are deposited in soil and covered by a frothy proteinaceous material produced in the female accessory glands. An acid-labile conjugate of 3,4dihydroxybenzoic acid was extracted from the accessory glands, and unconjugated 3,4-dihydroxybenzoic acid was obtained from the egg pods shortly after their deposition, indicating that it may be involved in sclerotization of the material (Hopkins et al., 1999). Some moths, such as saturniid silkmoths (Actias selene, Antheraea pernyi, Hyalophora cecropia, H. gloveri, and Samia cynthia), produce silk cocoons that are sclerotized after spinning. The silks are white when spun and remain white as long as they are in a dry atmosphere: When exposed to humid conditions a sclerotization process is initiated and the cocoons become brown. Glucosides of gentisic acid (23) and 3-hydroxyanthranilic acid (24) have been obtained from newly spun silks, and b-glucosidases and diphenoloxidases are present, indicating that silk sclerotization follow a pattern similar to that of the other sclerotization processes (Brunet and Coles, 1974). Manthey et al. (1992) reported that oxidation of 3-hydroxyanthranilic acid in the presence of proteins leads to formation of adducts between 3-hydroxyanthranilic acid and tyrosine residues in the proteins, and such adducts were isolated from cocoons of H. gloveri and S. cynthia. It appears that the adducts are formed via a radical– radical coupling mechanism. The newly spun silk of the Japanese giant silkmoth, Dictyoploca japonica contains a diphenoloxidase and a catecholic derivative, which was tentatively identified as N-(3,4-dihydroxyphenyllactyl)DOPA (25) (Kawasaki and Sato, 1985). It is apparently not glycosylated and no glucosidase activity was found in the silk. The presence of two catecholic groups in this sclerotization precursor may make it a more effective cross-linking agent than precursors with only a single catechol group.

4.4.4. Cuticular Enzymes and Sclerotization The study of cuticular enzymes has to a large extent been concerned with characterization of enzymes assumed to play a role in cuticular sclerotization,

mainly those involved in catechol oxidation and quinone isomerization, and there has been a tendency to neglect the possible presence of other types of enzymes that may play a role in cuticular stabilization. Catechol oxidation is an important step in sclerotization, wound healing, and immune responses in insects, and it is often difficult to decide whether a given cuticular phenoloxidase activity is of importance for sclerotization or whether its main role is to take part in defense reactions. It is therefore necessary to be extremely careful in interpreting the observations reported for cuticular enzymes. 4.4.4.1. ortho-Diphenoloxidases

After being transferred from the epidermal cells into the cuticle the catecholic sclerotization precursors may encounter various enzymes, such as o-diphenoloxidases, laccases, and peroxidases, capable of oxidizing them to quinones, but the relative roles of these activities are still uncertain. Insects contain inactive proenzymes for o-diphenoloxidases both in hemolymph and in the cuticle, and the proenzymes can be activated by a process initiated by wounds or the presence of small amounts of microbial cell wall components and involving limited proteolysis (Ashida and Dohke, 1980; Ashida and Brey, 1995). The o-diphenoloxidases can oxidize a wide range of o-diphenols, but not p-diphenols; they possess low hydroxylating activity towards monophenols, such as tyrosine and tyramine; and they are copper-containing enzymes readily inhibited by thioureas and Na-diethyldithiocarbamate. They have been isolated and characterized from both soft, nonsclerotizing cuticles, such as larval cuticle of Bombyx mori (Ashida and Brey, 1995; Asano and Ashida, 2001a), and from blowfly larval cuticles (Barrett, 1987a, 1987b, 1991) and sclerotizing pupal cuticle of Manduca sexta (Aso et al., 1984; Morgan et al., 1990). The amino acid sequences of o-diphenoloxidases from various insect species have been deduced from the corresponding DNA sequences (Fujimoto et al., 1995; Hall et al., 1995; Kawabata et al., 1995). The insect o-diphenoloxidases differ from diphenoloxidases (tyrosinases) from other organisms with regard to both substrate specificity and amino acid sequence (Sugumaran, 1998; Chase et al., 2000). According to the established sequences of insect prophenoloxidase genes, their gene products do not possess an N-terminal signal peptide sequence (Sugumaran, 1998), which is in contrast to what is commonly observed for proteins destined for export from the cells. The silk moth B. mori has genes for two o-diphenoloxidase proenzymes, and in the larvae the products of both genes are present in both

Cuticular Sclerotization and Tanning

hemolymph and cuticle, the only difference between the cuticular and hemolymphal forms being that several methionine residues are intact in the hemolymphal proenzymes, and they are oxidized to methionine sulfoxides in the cuticular proenzymes. The activated cuticular enzymes have nearly the same substrate specificity as the hemolymph enzymes; the main difference between the proenzymes is that in contrast to the unmodified hemolymph form the oxidized cuticular form cannot be transported across the epidermal cell layer, indicating that the epidermal transport of the proenzymes is a one-way traffic, from hemolymph to cuticle (Asano and Ashida, 2001a, 2001b). The enzymatic properties of the diphenoloxidases purified from hemolymph and from pharate pupal cuticle of M. sexta are very similar, suggesting a close relationship between the enzymes (Aso et al., 1985; Morgan et al., 1990), and it seems probable that they, like the B. mori enzymes, are derived from the same gene(s). When activated, both cuticular and hemolymph o-diphenoloxidases are very sticky i.e., they tend to aggregate and stick to any available surface and macromolecule, which will hinder diffusion of the active enzymes from the site where they became activated. The ultrastructural localization of o-diphenoloxidase activity in the larval cuticle of the blowfly Lucilia cuprina was described by Binnington and Barrett (1988) who observed activity in the epicuticular filaments. Activity was also observed in the procuticle, but only when the cuticle had been damaged beforehand, and the activity was limited to the close neighborhood of the wound, indicating that wounding is needed to activate the enzyme, and that the active enzyme remains in the vicinity of the wound. The prophenoloxidase in larval cuticle of B. mori was localized by immunocytochemical methods to the nonlamellate endocuticle, where it was randomly distributed, and to an orderly arrayed pattern in the lamellate endocuticle, but it appeared to be absent from the cuticulin layer and from the epidermal cells (Ashida and Brey, 1995). The role of the various cuticular o-diphenoloxidases in cuticular sclerotization is problematic; their presence as inactive proenzymes, which have to be activated, their close relationship to the hemolymphal phenoloxidases, and their abundance in nonsclerotizing cuticle indicate that their role is to take part in defense against wounding and microorganisms and not to be involved in sclerotization. It is, however, difficult to state with certainty that they do not play some role in sclerotization.

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4.4.4.2. Laccases

Laccase-type phenoloxidases have been reported to be present in dipteran larval cuticles shortly before and during puparium sclerotization, such as Drosophila virilis (Yamazaki, 1969), D. melanogaster (Sugumaran et al., 1992), Calliphora vicina (Barrett and Andersen, 1981), Sarcophaga bullata (Barrett, 1987a), and L. cuprina (Barrett, 1987b), and such enzymes have also been described from pupal cuticles of B. mori (Yamazaki, 1972) and M. sexta (Thomas et al., 1989) as well as from adult cuticle of the locust Schistocerca gregaria (Andersen, 1978). The nucleotide sequences for two laccase genes from M. sexta and a laccase gene from the mosquito Anopheles gambiae have recently been deposited in the GenBank, and the accession numbers for the corresponding proteins are: AAN1706, AAN1707, and AAN17505, respectively. The insect laccases are structurally related to laccases of plant or fungal origin. In contrast to the insect diphenoloxidases the laccase gene products contain a typical signal peptide sequence, indicating that the enzymes are secreted into the extracellular space. In larval cuticles of D. virilis (Yamazaki, 1969) and L. cuprina (Binnington and Barrett, 1988) laccase activity makes its appearance shortly before pupariation. In both species the enzyme activity decreases gradually as puparial sclerotization progresses. Laccase activity can be demonstrated a few days before ecdysis in pharate cuticle of adult locusts, S. gregaria; it remains at high levels for at least 2 weeks after ecdysis, and activity has also been demonstrated in nymphal exuviae, indicating that the locust enzyme is not inactivated by sclerotization (S.O. Andersen, unpublished data). Laccases are active towards a broad spectrum of o- and p-diphenols: NBAD and NADA are among the best o-phenolic substrates tested, and methylhydroquinone is the best p-diphenolic substrate. Insect laccases are not inhibited by compounds, such as thiourea, phenylthiourea, and Na-diethyldithiocarbamate, which are effective inhibitors of o-diphenoloxidases, but they are inhibited by carbon monoxide and millimolar concentrations of fluorides, cyanides, and azides (Yamazaki, 1972; Andersen, 1978; Barrett and Andersen, 1981; Barrett, 1987a). The laccases are resistant towards treatments inactivating many enzyme activities; the S. gregaria laccase remains active after blocking available amino and phenolic groups by dinitrophenylation or dansylation, and it survives temperatures up to about 70
C, but it is inactivated by treatment with tetranitromethane, which nitrates tyrosine residues (Andersen, 1979b). The laccases

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appear to be firmly linked to the cuticular structure; typically they cannot be extracted by conventional protein extractants, but are readily extracted after limited tryptic digestion of the yet-unhardened cuticle (Yamazaki, 1972; Andersen, 1978). The enzyme was obtained from C. vicina larval cuticle by prolonged extraction at pH 8 without addition of any protease, but as latent protease activity is present in the cuticle the release of laccase from the cuticular residue may be due to proteolysis (Barrett and Andersen, 1981). The enzyme is not released by tryptic digestion of already sclerotized cuticle. The ultrastructural localization of laccase activity has been studied in the L. cuprina larval cuticle (Binnington and Barrett, 1988) and enzyme activity was observed in the inner epicuticle of late third instar larvae (about to pupariate), but not in epicuticle of younger larvae. The laccase activity in L. cuprina larval cuticle could be demonstrated without prior activation, in contrast to the cuticular o-diphenoloxidases, indicating that the laccase is not deposited as an inactive precursor in this insect, and neither is an inactive proenzyme likely to be present in pharate locust cuticle since enzyme activity could be demonstrated without any activating treatment. A pro-laccase has been purified and partially characterized from cuticle of newly pupated pupae of B. mori (Ashida and Yamazaki, 1990). The inactive pro-laccase could be activated by treatment with various proteolytic enzymes, and the substrate specificities of the laccase variants obtained depended upon the protease used for activation. 4.4.4.3. Cuticular Peroxidases

Several different routes for the oxidation of catechols to o-quinones may be of advantage for an insect, especially if they can be regulated independently in various cuticular regions, and peroxidase activity may provide such an alternative route. Peroxidase activity has been demonstrated by histochemical methods in proleg spines of Calpodes ethlius larvae (Locke, 1969) and in larval and pupal cuticle of Galleria mellonella and Protophormia terraenovae (Grossmu¨ ller and Messner, 1978; Messner and Janda, 1991; Messner and Kerstan, 1991), and such activity is also observed intracellularly in different cell types in insects. It is not known whether the cuticular peroxidase activities are identical to the intracellular enzymes, as the cuticular activity has never been properly characterized. Proteins can be cross-linked by means of the peroxidase system, and it has been suggested that the enzyme could be involved in cuticular sclerotization (Hasson and Sugumaran, 1987). A peroxidase is likely to be

involved in the cross-linking of the rubberlike elastic cuticular protein resilin (Andersen, 1966; Coles, 1966). This cuticular protein is cross-linked by oxidative coupling of tyrosine residues during its extracellular deposition, and the tyrosine radicals needed for the coupling may be formed by a peroxidase catalyzed oxidation process. Peroxidases can also oxidize catechols to semiquinone radicals, two of which readily dismutate to form an o-quinone and a catechol. The enzyme needs hydrogen peroxide as one of its substrates, and Candy (1979) reported that locust cuticle contains a glucose oxidase activity, which oxidizes glucose to d-gluconate with concomitant production of hydrogen peroxide. It was suggested that the hydrogen peroxide produced may participate in sclerotization reactions. Candy (1979) also reported the presence in locust cuticle of several other enzymes involved in hydrogen peroxide metabolism, such as peroxidase, catalase, and superoxide dismutase. Peroxidase activity in solid cuticle may be involved in the production of dityrosine cross-links and in oxidizing catechols to quinones for sclerotization. Dityrosine has so far not been demonstrated in sclerotized cuticle from insects, but dityrosine as well as brominated dityrosines have been obtained from the hardened exocuticle of the crab Cancer pagurus (Welinder et al., 1976). The eggshells of D. melanogaster are stabilized by formation of dityrosine cross-links between the protein chains (Petri et al., 1976; Mindrinos et al., 1980), and the hardening of Aedes aegypti egg chorion includes both peroxidase-mediated protein cross-linking through dityrosine formation and diphenoloxidasecatalyzed chorion melanization (Li et al., 1996). The hydrogen peroxide necessary for dityrosine formation in A. aegypti chorion is produced by an enzymatic process by which NADH is oxidized with concomitant reduction of molecular oxygen to hydrogen peroxide. The necessary supply of NADH for this process is provided by enzyme-catalyzed oxidation of malate coupled to reduction of NADþ (Han et al., 2000a, 2000b). It is unknown whether a similar system for providing hydrogen peroxide operates during sclerotization of insect cuticles. 4.4.4.4. ortho-Quinones and para-Quinone Methides

The three types of oxidases, o-diphenoloxidases, laccases, and peroxidases, can all oxidize NADA and NBAD to their respective o-quinones. The o-quinones are reactive compounds, they can spontaneously form adducts by reaction with available nucleophilic groups, and they can serve as substrates for other cuticular enzymes, o-quinone and p-quinone

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methide isomerases, catalyzing isomerization to p-quinone methides, which also react readily with nucleophilic compounds. In the traditional sclerotization scheme, suggested by Pryor (1940a, 1940b), o-quinones were supposed to react preferably with e-amino groups from lysine residues, but solid state nuclear magnetic resonance (NMR) studies have shown that the imidazole group in histidine residues is the preferred cuticular target for reaction with quinones (Schaefer et al., 1987; Christensen et al., 1991). Incubation of blowfly (Sarcophaga bullata) larval cuticle with NADA and N-acetylcysteine resulted in formation of an adduct (26) where the sulfur atom is linked to the 5-position of the NADA moiety (Sugumaran et al., 1989a), indicating that SH-groups are good acceptors for oxidized NADA (Figure 5). Electrochemical oxidation of dopamine to dopamine quinone in the presence of N-acetylcysteine gave a mixture of C-5 and C-2 (27) monoadducts together with some of the disubstituted product, 2,5-S,S0 -di(N-acetylcysteinyl)dopamine (28) (Xu et al., 1996a; Huang et al., 1998). Since the monoadducts are more readily oxidized to quinones than is the parent catechol, a monoadduct formed between an oxidized catechol and a protein-linked cysteine will be more prone than a free catechol to be reoxidized to quinone, which can react with a cysteine residue in neighboring protein chain to form a covalent cross-link between the proteins. Thus cysteine–catechol based cross-links are possible,

Figure 5 Adducts formed by reaction of N-acetylcysteine with the o-quinones of NADA and dopamine. 26, 5-S-(N-acetylcysteinyl)-N-acetyldopamine; 27, 2-S-(N-acetylcysteinyl)-dopamine; 28, 2,5-S,S0 -di-(N-acetylcysteinyl)-dopamine.

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but they appear not to play an important role in cross-linking cuticular proteins since these proteins typically contain neither cysteine nor cystine. Among all the cuticular proteins that have been sequenced so far, only a few contain residues of sulfur-containing amino acids, and it seems unlikely that sulfur can have an important role in sclerotization. Perhaps the scarcity of cystine and cysteine in cuticular proteins is related to the readiness with which they react with o-quinones. When locust cuticle is incubated with NADA and benzenesulfinic acid, the oxidized NADA is trapped by adduct formation with the sulfinic acid, and before all sulfinic acid has been consumed by adduct formation no o-quinone is available for reaction with cuticular proteins or isomerization and further metabolism to polymeric compounds (S.O. Andersen, unpublished data). The presence of significant amounts of free SH-groups in the cuticular matrix proteins could in a similar way delay further metabolism of the o-quinones and thereby cause suboptimal sclerotization. Electrochemical oxidation of NADA to NADAquinone in the presence of N-acetylhistidine gave monoadducts (Figure 6), where a nitrogen atom in the imidazole ring is linked to either the C-6 (29) or the C-2 ring position (30) in NADA, the C-6 position being the preferred position (Xu et al., 1996b). Electrochemical characterization of the C-6 and C-2 N-acetylhistidine-NADA adducts showed that both adducts are more difficult to oxidize than NADA itself (Xu et al., 1996b), indicating that formation of adducts involving two N-acetylhistidine residues linked to the same NADA residue is not very likely. Oxidation of a mixture of NADA plus N-acetylhistidine by means of larval cuticle from H. cecropia gave a mixture of products, and adduct formation to the C-6 ring position as well as to the b-position of the side chain (31) was observed (Figure 6) (Andersen et al., 1991, 1992c). The formation of a side chain adduct indicates activation of the b-position, probably due to formation of the p-quinone methide, demonstrating that cuticular oxidation of catechols is more complex than electrochemical oxidation. From acid hydrolysates of M. sexta pupal cuticle, adducts have been obtained where histidine is linked to either the C-6 ring position or the b-position of dopamine or to the corresponding positions in DOPET, demonstrating that adduct formation to histidine residues occurs during in vivo sclerotization (Xu et al., 1997; Kerwin et al., 1999), and confirming that DOPET has a role as a sclerotization precursor. Direct evidence that covalent bonds are formed between acyldopamines and

156 Cuticular Sclerotization and Tanning

Figure 6 Adducts formed by reaction of N-acetylhistidine with NADA-o-quinone or NADA-p-quinone methide. 29, 6-(N-acetylhistidyl)-N-acetyldopamine; 30, 2-(N-acetylhistidyl)-N-acetyldopamine; 31, 7-(N-acetylhistidyl)-N-acetyldopamine.

histidine residues during sclerotization had previously been obtained by solid state NMR studies, utilizing incorporation of isotopically labeled dopamine, histidine, and b-alanine into sclerotizing pupal cuticle of M. sexta (Schaefer et al., 1987; Christensen et al., 1991). The spectra demonstrated the presence of bonds between nitrogen atoms in the imidazole ring of histidine and ring positions and b-position of the side chain of dopamine. The formation of covalent bonds involving the amino group of b-alanine and the e-amino group of lysine was also indicated. Furthermore, catecholamine-containing proteins have been purified and partially characterized from sclerotizing M. sexta pupal cuticle, and NBANE was released from these proteins on mild acid hydrolysis, indicating the presence of a bond between the b-position of the side chain of NBAD and some amino acid residues in the proteins (Okot-Kotber et al., 1996). Incubation of larval cuticle of H. cecropia with NADA together with compounds containing a free amino group, a-N-acetyllysine or b-alanine, resulted in the formation of a number of products (Figure 7), and adducts were identified with the amino groups linked to the 6-position of the ring in NADA (32). These adducts have a quinoid structure, indicating that the primary catecholic product is readily oxidized, in contrast to the adducts formed to the b-position of the side chain (33), which are stable in their catecholic form (Andersen et al., 1992b). Adduct formation to either histidine and lysine residues during cuticular oxidation of NADA appears to be so much slower than adduct formation to cysteine that a significant fraction of the o-quinones obtained from NADA or NBAD will be available for the isomerase catalyzing further metabolism of the quinones. Prolonged incubation of NADA and

Figure 7 Adducts formed by cuticular oxidation of a mixture of NADA and a-N-acetyllysine.

a-N-acetyllysine together with cuticle resulted in the formation of 4-phenylphenoxazin-2-ones (34), which are composed of three NADA residues joined to one a-N-acetyllysine residue (Peter et al., 1992).

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The structure of one of the products formed during incubation of insect cuticles with NADA plus N-acetylamino acids indicated that o-quinone of NADA can react with water to form N-acetyl-3,4,6-trihydroxyphenylethylamine (6-hydroxy-NADA), which then couples oxidatively with another NADA residue to give the dimeric compound (35) (Figure 8) (Andersen et al., 1992a). Corresponding derivatives indicating formation of 6-hydroxy-NADA have so far not been reported from naturally sclerotized cuticle, and its formation in vitro may be due to the large excess of water in incubation experiments, in contrast to the relatively low water content in sclerotizing cuticle (30–40% of the cuticular wet weight). The incubation of cuticle together with NADA resulted also in the production of small amounts of a product (36) consisting of two NADA-quinones linked together via their 6-positions (Andersen et al., 1992a). A corresponding 2,60 -linked dimer of NADA was suggested as intermediate in the formation of the above mentioned 4-phenylphenoxazin-2-ones (34) (Peter et al., 1992). The side chain b-position in the p-quinone methides reacts readily with water, and the lack of stereospecificity in the reaction indicates that the addition of water is nonenzymatic (Peter and Vaupel, 1985). The relative high yields of NANE and NBANE obtained by extraction of various cuticles are probably due to both this reaction and hydrolysis of labile adducts formed with other nucleophiles, such as hydroxyl groups in chitin (see Section 4.4.3.2.2). The enzyme responsible for quinone isomerization has been partially characterized from larval cuticles from H. cecropia (Andersen, 1989c) and the flesh fly S. bullata (Sugumaran, 1987; Saul and Sugumaran, 1988), both cuticles belonging to the soft, pliant type. The enzyme is also present in the hemolymph of S. bullata (Saul and Sugumaran, 1989a, 1990), where it participates in defense reactions. The enzyme has thus been described from both sclerotizing and nonsclerotizing cuticles and from insect hemolymph, leading to the question whether the same enzyme is involved in both defense reactions and sclerotization. It would be worthwhile

Figure 8 Products obtained by cuticular oxidation of NADA.

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to study the temporal and regional distribution of the enzyme in various insect systems. 4.4.4.5. Dehydro-NADA and Dehydro-NBAD

The p-quinone methide formed by isomerization of NADA-o-quinone can be further isomerized to dehydro-NADA, a NADA derivative carrying a double bond between the a- and b-carbon atoms in the side chain. The enzyme responsible for this isomerization has been called N-acetyldopamine quinone methide/1,2-dehydro-N-acetyldopamine tautomerase (Saul and Sugumaran, 1989c). The activity has been reported to be present in larval cuticle of S. bullata (Saul and Sugumaran, 1989b, 1989c) and D. melanogaster (Sugumaran et al., 1992). A related enzyme activity catalyzing the isomerization of NBAD quinone methide to dehydro-NBAD has been demonstrated in extracts of C. vicina larval cuticle (Ricketts and Sugumaran, 1994). It is probably the same enzyme which leads to the formation of dehydro-NADA and dehydro-NBAD. Locust cuticle catalyzes formation of dehydro-NADA from NADA, whereas NBAD is a poor substrate for the locust enzyme (Andersen, 1989d). It appears that several cuticles, which are sclerotized by mixtures of NADA and NBAD, readily convert NADA to the dehydro-derivative, while conversion of NBAD to dehydro-NBAD only occurs to a minor extent, if at all (Andersen, 1989d; Andersen et al., 1996). It is uncertain whether formation of dehydroNADA in locust cuticle occurs only via NADAquinone methide or whether it can also be formed directly from NADA by a route circumventing quinone formation. Inhibition of oxidation of NADA to NADA-quinone resulted in accumulation of dehydro-NADA in locust cuticle, and since it seemed unlikely that accumulation of a product can be caused by decreased production of its precursors, the presence of a specific ‘‘NADA-desaturase’’ was suggested (Andersen and Roepstorff, 1982; Andersen et al., 1996). The dehydro-NADA formed during cuticular sclerotization can be oxidized by both the o-diphenoloxidases and laccases present in cuticle, and the resulting unsaturated quinones react spontaneously with other catechols to give substituted dihydroxyphenyl-dihydrobenzodioxines (XIII) (Andersen and Roepstorff, 1982; Sugumaran et al., 1988). It has not been established whether the unsaturated quinones will also react with nucleophilic residues in cuticular proteins. Locust cuticle will catalyze in vitro formation of adducts between dehydro-NADA and catechols, and dihydroxyphenyl-dihydrobenzodioxine derivatives, corresponding to those extracted from naturally sclerotized cuticle, can be formed in this

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way (Andersen and Roepstorff, 1982; Andersen, 1985). It appears that the presence of dihydroxyphenyl-benzodioxine derivatives in sclerotized cuticles can be taken as an indication for the presence of a dehydro-NADA forming activity. It has been suggested that the cuticular diphenoloxidases, isomerases, and tautomerases occur together in large enzyme complexes, enabling one enzyme to deliver its products directly to the following enzyme for further processing (Andersen et al., 1996; Sugumaran, 1998). The evidence for such complexes is insufficient, but the observation that during in vitro incubation with NADA some cuticles convert NADA into benzodioxine dimers with very little formation of NANE, while other cuticles preferably produce NANE, may indicate that the former cuticles contain enzyme complexes with tight coupling between the individual enzymes, so the intermediates will have little chance of reacting with water or the surrounding proteins before they are used as substrates for the next enzyme in the sequence. The sclerotization of some types of cuticle may thus be dominated by reactions involving the dehydro-NADA o-quinone, while sclerotization of other types of cuticle is dominated by reactions between the initially generated quinones and the matrix proteins. The final outcome of the sclerotization process probably depends upon a carefully controlled balance between various enzyme activities and available substrates, a balance that differs between the local cuticular regions according to their mechanical properties. It would be interesting to have quantitative determinations of the relevant enzyme activities in different types of cuticle, and to be able to specifically inhibit the individual enzymes participating in sclerotization to study their individual roles in the total process. Quantitative enzyme activity determinations on pieces of insoluble materials are problematic, due both to slow diffusion of substrates into, and products out of, the pieces and because it is difficult to obtain homogeneous samples for analysis. However, it should be possible to obtain relative values for the activities to be used for comparison of various cuticular regions. 4.4.4.6. Various Catechol Derivatives Obtained from Cuticles

An indication of the quantitative role of the possible sclerotization reaction pathways may be obtained by identification and quantification of the low molecular weight catecholic compounds, which can be extracted from sclerotized cuticles after more or less extensive degradation of the cuticular material. The structures of such compounds may suggest whether

they are likely to be products derived from the sclerotizing precursors and perhaps give some indication of the reactions responsible for their generation. The compounds extracted from sclerotized cuticle could represent unused sclerotization precursors, intermediates in the sclerotization process, by-products from the process, and degradation products of protein-bound cross-links and polymers. It is also possible that the extracted compounds have their own distinct functions in the cuticle, quite unrelated to sclerotization; i.e., catechols can thus be precursors for pigments, such as papiliochromes (Umebachi, 1993), or they can function as antioxidants protecting the epicuticular lipids from autoxidation (Atkinson et al., 1973a). 3,4-Dihydroxyphenylacetic acid, which is present in the solid cuticle of many beetle species (Andersen, 1975), could serve the latter purpose, as it apparently takes no part in the sclerotization process (Barrett, 1984, 1990). A detailed study of the formation and fate of the various cuticular compounds is necessary for deciding whether they are related to the sclerotization process. The mixture of compounds obtained from cuticles by the use of mild extraction methods, such as extraction in distilled water or neutral salt solutions at moderate temperatures, contains compounds that are less modified than those obtained by acidic extraction at elevated temperatures, but a critical and careful interpretation of their structures will be necessary in all cases. Extraction with dilute acids tends to give higher yields of catechols than extraction with water, but the compounds identified in the extracts are often the same (Atkinson et al., 1973b). The higher yield obtained with acids may be due partly to swelling of the cuticular material at low pH values, resulting in easier liberation of trapped compounds, and partly to hydrolysis of acid-labile bonds. Typical extraction products are the sclerotization precursors, NADA and NBAD, and their hydroxylated derivatives, NANE and NBANE, and they can also occur as O-glucosyl derivatives. The O-glycoside linkage is acid-labile and may not survive prolonged extraction, and the b-hydroxyl group in NANE and NBANE may either be produced by reaction of the p-quinone methides of NADA and NBAD with water, or can result from the hydrolysis of the products formed by reaction of a p-quinone methide with cuticular proteins and chitin. 3,4-Dihydroxybenzoic acid (18) and 3,4-dihydroxybenzaldehyde have been extracted from several types of sclerotized cuticle; they are probably formed by extensive oxidative degradation of the side chain of the sclerotizing precursors, NADA and NBAD. The precise reaction pathway for their

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Figure 10 43, Trimeric dihydrobenzodioxine derivative from sclerotized locust cuticle.

Figure 9 Ketocatechols obtained by mild acid hydrolysis of sclerotized insect cuticle. 37, arterenone; 38, 3,4-dihydroxyphenylketoethanol (DOPKET); 39, 3,4-dihydroxyphenylglyoxal; 40, 3,4-dihydroxyphenylglyoxylic acid; 41, N-acetylarterenone; 42, 3,4-dihydroxyphenylketoethylacetate.

formation is not known, but their common occurrence in sclerotized cuticle indicates that the intracuticular environment is highly oxidative during the sclerotization process. The presence of dopamine and norepinephrine in cuticular extracts may be due to deacylation of the N-acylated forms, or they may have been transferred directly from epidermal cells to cuticle, either by active transport across the apical cell membrane or by passive leakage through the cell membrane. The black cuticle of the fruitfly mutants black and ebony contains elevated levels of dopamine (Wright, 1987), which probably have been transferred to the cuticle in the nonacylated state. Quite large amounts of ketocatechols (Figure 9), such as arterenone (37), DOPKET (38), and N-acetylarterenone (41), as well as 3,4-dihydroxyphenylglyoxal (39), 3,4-dihydroxyglyoxylic acid (40), and 3,4-dihydroxyphenylketoethylacetate (42), can be obtained from sclerotized cuticle by treatment with dilute acids (Andersen, 1970, 1971; Andersen and Barrett, 1971; Andersen and Roepstorff, 1978). The yields of ketocatechols can amount to several percent of the cuticular dry weight (Andersen, 1975; Barrett, 1977), and the type of ketocatechols released depends upon the exact conditions of hydrolysis. It has been argued that they are all

degradation products of a common precursor in the cuticle (Andersen, 1971). More complex catechol derivatives can be obtained by extracting sclerotized cuticle at relatively mild conditions, such as concentrated formic acid at ambient temperature or boiling dilute acetic acid. From such extracts a number of dimeric compounds of the dihydroxyphenyl-dihydrobenzodioxine type were isolated and identified (Andersen and Roepstorff, 1981; Roepstorff and Andersen, 1981), and later a trimeric compound (44) (Figure 10) was obtained by formic acid extraction of sclerotized locust cuticle (Andersen et al., 1992c). Ketocatechols are readily formed when such dimers and oligomers are hydrolyzed with acid (Andersen and Roepstorff, 1981), but the extractable material accounts for only a minor fraction of ketocatechols formed by acid hydrolysis of sclerotized cuticle, and the main fraction of ketocatechols obtainable from cuticle is presumably derived from catecholic material covalently linked to the cuticular proteins and chitin. Since the various dimers can be formed in vitro by reaction between oxidized dehydroNADA and catechols, it is likely that oxidized dehydro-NADA will react with catechols (NADA or NBAD) linked to cuticular proteins to form protein-linked dimers and higher oligomers.

4.4.5. Control of Sclerotization Sclerotization of insect cuticles varies regionally both with respect to time of initiation, intensity, duration, and balance between NADA- and NBAD-sclerotization, indicating that the process must be controlled locally to give the optimal result. 4.4.5.1. Pre-Ecdysial Sclerotization

In some cuticular regions sclerotization starts in the pharate stage, and these regions attain their final size and shape during the pre-ecdysial deposition of cuticular materials (Cottrell, 1964). Pre-ecdysial

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sclerotization may be limited to small, local regions, such as mandibles and spines, or larger regions of cuticle, covering thorax, head, and legs, may be stabilized before ecdysis. In the presclerotized regions sclerotization continues after ecdysis, whereby the already stiffened material is further strengthened. Pre-ecdysial sclerotization has been observed in several cuticular regions in adult honeybees (Apis mellifera adansonii) (Andersen et al., 1981), and the matrix proteins in these regions resist extraction with solvents that do not degrade the cuticle. Acid hydrolysis of pre-ecdysial sclerotized honeybee pharate cuticle yielded significant amounts of ketocatechols, and still more was obtained by hydrolysis of the corresponding cuticular regions from mature worker bees where endocuticle deposition was complete, indicating that the same type of sclerotization is involved in both pre- and post-ecdysial stabilization. The wings were the only body region where no pre-ecdysial sclerotization was observed. Their sclerotization occurred rapidly after ecdysis and was nearly complete when the bees left the cell in which they had pupated, confirming that insect wings are not stabilized until they have been expanded to their proper size after emergence. 4.4.5.2. Post-Ecdysial Sclerotization

Many regions of the cuticle are often expanded to a new and bigger size after the insect has emerged from the old cuticle (Cottrell, 1964). In some insects the period from ecdysis to fully expanded cuticle can be rather prolonged, for instance in flies where the newly emerged adult has to dig its way through the substratum in which it pupariated, but many insects can start cuticular expansion as soon as they have escaped from the exuvium. Release of the neurohormone bursicon from the central nervous system is a signal for initiating general sclerotization after ecdysis. Bursicon has a pronounced influence on the activities of the epidermal cells. It has been reported that lack of bursicon results in the failure of endocuticle deposition as well as melanin production and sclerotization of the cuticle (Fraenkel and Hsiao, 1965; Fogal and Fraenkel, 1969), and it appears that bursicon is involved in the control of tyrosine hydroxylation to DOPA (Seligman et al., 1969). In some insects sclerotization stops when the preecdysially deposited cuticle has become sclerotized to form exocuticle, although deposition of endocuticle continues for several days, resulting in a sclerotized exocuticle and a nonsclerotized endocuticle. In other insects cuticular sclerotization continues during endocuticle deposition with the result that both exo- and endocuticle become sclerotized, but not to the same degree. Sclerotization of femur cuticle in

adult locusts (Schistocerca gregaria) continues for at least 12 days after ecdysis, and both exo- and endocuticle are sclerotized (Andersen and Barrett, 1971), in contrast to sclerotization of the femur cuticle of fifth instar nymphs of the same species which lasts for only 1 day, and deposition of unsclerotized endocuticle continues for about 4–5 days (Andersen, 1973). Accordingly, the endocuticular proteins are readily extractable from femurs of mature nymphs, but little protein can be extracted from the femurs of mature adult locusts. The difference in duration of sclerotization in locust nymphs and adults is probably related to the different fate of these two types of cuticle. The nymphal cuticle will to a large extent be degraded in preparation for the next ecdysis, and sclerotized cuticle is more resistant to enzymatic degradation than nonsclerotized cuticle. The adult cuticle has to last for the rest of the life of the animal, and there is no apparent advantage in having an easily degraded endocuticle. The leg cuticle of adult locusts is also exposed to stronger mechanical forces than the cuticle of nymphal legs, and it may therefore be an advantage for adult locusts to have both layers of the leg cuticle stabilized, but not to the same degree. A similar sclerotization difference between adult cuticle and cuticle in younger instars is probably present in other insect species. It has not yet been established how the duration of the post-ecdysial sclerotization is regulated. 4.4.5.3. Puparial Sclerotization

During puparium formation in the higher Diptera the soft, pliable cuticle of the last larval instar is modified to a hard and nondeformable material, its function being mainly to protect the animal during pupal and adult development and not to allow movements. Pronounced regional differentiation in cuticular sclerotization is apparently not essential for puparia. The chemical processes of puparium sclerotization are very similar to those involved in the sclerotization of adult cuticle, but the two systems differ in the way they are controlled (Sekeris, 1991). Puparium sclerotization begins with an increase in ecdysteroid titer, which induces the expression of the enzyme DOPA decarboxylase, catalyzing the decarboxylation of DOPA to dopamine. The latter is then acylated to the sclerotization precursors, NADA and NBAD, a process catalyzed by the enzymes acetyl transferase and b-alanyl transferase, respectively. The precursors are oxidized to o-quinones, which are then isomerized to p-quinone methides which in turn may be converted to the corresponding dehydro-derivatives (Saul and Sugumaran, 1989a; Sugumaran et al., 1992). The flies Musca autumnalis and M. fergusoni harden

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their puparia by the deposition of calcium and magnesium phosphates and not by phenolic sclerotization (Gilby and McKellar, 1976; Darlington et al., 1983). 4.4.5.4. Transport of Sclerotization Precursors to the Cuticle

Cuticle of actively moving insects is constantly exposed to external forces which vary in intensity and direction, and to keep the resulting local deformations of the cuticle within functionally acceptable ranges, the mechanical properties of each of the cuticular regions will probably have to be rather precisely regulated, suggesting that the degree of sclerotization of the individual regions is controlled by local mechanisms. In all likelihood such local sclerotization control resides in the underlying epidermal cells. It could operate by regulating the supply of sclerotization precursors to the cuticle, for instance by controlling local synthesis of precursors and/or their uptake from hemolymph into epidermis or by controlling the transport of precursors from epidermal cells to the cuticular matrix where they will be exposed to the enzymes converting them to sclerotization agents. Precursors for sclerotization can be synthesized in the epidermal cells, but it is likely that they can also be produced by other cell types, such as hemocytes and/or fat body cells. Precursors, such as dopamine, NADA, and NBAD, injected into the hemocoel shortly before or during cuticular sclerotization, are rapidly taken up by the epidermal cells and transported into the cuticle, whereas precursors injected several days before the start of sclerotization are mainly degraded or modified by glycosylation or phosphorylation. Such precursor conjugates may either remain in the animal and serve as a reserve pool of sclerotizing material, or they may be excreted via the Malphigian tubules by standard detoxification mechanism. The white pupa mutant of the Mediterranean fruit fly, Ceratitis capitata, fails to sclerotize the puparium, but develops normal larval and adult cuticles. The concentrations of the various catecholamines were very low in the mutant puparial cuticle compared to the wild-type strain, whereas the concentrations in the hemolymph of NADA, NBAD, and dopamine were about ten times higher in the mutant than in the wild-type, indicating that the mutant is defective in the system transporting catecholamines from hemolymph to puparial cuticle (Wappner et al., 1995). To control the transport of sclerotizing precursors into the cuticle in the right amounts at the right time the epidermal cells must possess specific transport systems both in the basolateral cell membrane to

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facilitate uptake from the hemolymph and in the apical cell membrane to control the transport of sclerotizing precursors from the cells into the cuticle. Active diphenoloxidases (laccases) are, at least in some insects, present in unsclerotized pharate cuticle before or at ecdysis (Andersen, 1972, 1979b, and unpublished data), and it appears unlikely that the precursors are transported into these cuticles before sclerotization is initiated following ecdysis. Radioactive NADA injected into fifth instar locust nymphs 1–2 days before ecdysis, when deposition of the pharate adult cuticle occurs, is partly incorporated into the old nymphal cuticle, soon to be discarded, partly retained to be incorporated into the new adult cuticle after ecdysis, and partly excreted as an O-glucoside (Andersen, 1974b and unpublished data). The labeled NADA, which after ecdysis was incorporated into the new cuticle, was in the meantime probably stored in the epidermal cells, but other locations, such as the fat body or hemocytes, cannot be disregarded, and the mechanism for transferring NADA into the cuticle is unexplored. It has been reported that proteins to which catechol derivatives are attached can be transported intact from the hemolymph to the cuticular structure to serve as combined matrix components and sclerotization precursors (Koeppe and Mills, 1972; Koeppe and Gilbert, 1974; Bailey et al., 1999), indicating that receptors able to recognize such proteins must be present in the basolateral membrane of the epidermal cells. 4.4.5.5. Balance between Cuticular Enzymes

The type of sclerotization occurring in the local regions will depend in part upon the relative amounts of the two precursor compunds, NADA and NBAD, imported from the epidermal cells, and partly upon the balance between the various enzyme activities in the cuticle. An attempt to study some of these questions was made 30 years ago (Andersen, 1974a, 1974b), but the means available at that time were insufficient to give conclusive answers. NADA labeled with tritium either on the aromatic ring or on the b-position of the side chain was used to study the extent to which the ring and the side chain were involved in adduct formation, based on the assumption that sclerotization via an o-quinone will result in tritium release from the ring and that release of tritium from the NADA side chain will occur without intermediate quinone formation. Different types of cuticle were found to differ in their ability to release tritium from the two positions; lightly colored, sclerotized cuticles preferentially released tritium from the side chain, and dark-brown cuticles

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released tritium preferentially from the ring system (Andersen, 1974a). It is now evident that tritium release from the b-position of the side chain will occur during formation of the p-quinone methide of NADA and will be a measure of the o-quinone isomerase activity present, and that tritium release from the aromatic ring will occur during adduct formation between an o-quinone and some nucleophile. This will be a measure of the fraction of available NADA that is oxidized by the cuticular diphenoloxidase, but escapes being isomerized to a p-quinone methide. Release of tritium from NADA specifically labeled as the a-position of the side chain can presumably be used for obtaining an estimate of the relative importance of the formation of dehydro-NADA during cuticular sclerotization (Andersen, 1991). Locust (S. gregaria) femur cuticle was found to release little tritium from the ring system of NADA, but significant and nearly equal amounts of tritium from the a- and b-positions of the NADA side chain (Andersen, 1974a and unpublished data), indicating that almost all NADA used for sclerotization by this insect is converted to 1,2-dehydro-NADA. Since NADA and NBAD appear to be treated differently during cuticular sclerotization, it will be necessary to use a- as well as b-tritiated forms of both NADA and NBAD to determine the quantitative importance of p-quinone methide sclerotization relative to sclerotization involving the dehydro-derivatives. Such determinations have, to the best of our knowledge, not been reported. 4.4.5.6. Intensity of Cuticular Sclerotization

It is difficult to obtain a useful measure of the degree of sclerotization of a given piece of cuticle, but due to its importance for the mechanical properties of cuticles, it would be an advantage to have some means for determining how much NADA and NBAD are incorporated during sclerotization. It is possible that solid state NMR studies can be used for the measurements (Schaefer et al., 1987; Christensen et al., 1991), but such techniques are not generally available. Attempts to determine the degree of sclerotization of various cuticular samples have been made by measuring the amounts of ketocatechols released by acid hydrolysis from cuticular samples (Andersen, 1974b, 1975; Barrett, 1977, 1980), but this method only determines that part of total sclerotization which is due to reactions involving activated dehydro-NADA and dehydro-NBAD. A useful measure for the quantitative role of NBAD in sclerotization might be obtained by determining the amounts of b-alanine released by acid hydrolysis; it appears that some of the amino groups in b-alanine take

part in the sclerotization process and cannot be released by hydrolysis (Christensen et al., 1991), but this will probably only represent a minor fraction of the total amount of b-alanine incorporated. Although analysis for ketocatechols as well as balanine cannot give absolute values for the types of sclerotization they represent, relative values can be useful for comparing different cuticular samples. Ketocatechols were not obtained from adult femur cuticle of the locust S. gregaria when the samples were taken just after the animals had emerged from their exuvium, as no sclerotization had yet occurred. Later, during maturation of the locusts, a steady increase in the yield of ketocatechols was observed, reaching a constant level after about 1 week, when the ketocatechol yield was about 0.2 mmol mg1 dry cuticle (Andersen and Barrett, 1971), indicating a close relationship between ketocatechol formation and sclerotization in locust cuticle. Determination of ketocatechol release from various types of cuticle from mature locusts (S. gregaria) showed that all cuticular types yielded some ketocatechols upon hydrolysis: from 10–12 days old locusts the lowest yields were obtained from abdominal intersegmental membranes (0.20% of the dry weight), abdominal sclerites gave 0.38% of the dry weight, and the highest values were obtained from the dorsal mesothorax (5.25% of the dry weight) and the mandibles (3.36% of the dry weight) (Andersen, 1974b), in agreement with the expectation that the regions where strength and hardness are essential for proper function are also the regions giving the highest yields of ketocatechols. Schistocerca gregaria mature nymphal cuticle gave much lower amounts of ketocatechols than corresponding samples from mature adults. The only exception was the nymphal mandibles; 3.74% of their dry weight was recovered as ketocatechols, comparable to what was obtained from the adult mandibles (Andersen, 1974b). The relative low yield of ketocatechols obtained from nymphal cuticle are probably related to the nymphs weighing less than adults, their leg cuticle being exposed to smaller forces during walking and jumping than the legs of adults, and the nymphal wings and thorax cuticle not being exposed to deforming forces comparable to those for which adult cuticle is exposed to during flight. Quantitative ketocatechol determinations have been performed on a few other insect species, such as the beetles Tenebrio molitor and Pachynoda epphipiata (Andersen, 1975). Various parts of the exuviae of a cicada, Tibicen pruinosa, have also been analyzed (Barrett, 1977), and most regions

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gave ketocatechol values between 6% and 7% of the exuvial dry weight. The exuvial cuticle from the cicada compound eyes gave a ketocatechol yield (11.8% of the dry weight) that was much higher than that obtained from any of the other cuticular regions. The high values obtained from exuviae agree with the notion that sclerotization is generally more pronounced in exocuticle than in endocuticle. The very high values obtained from exuvial cornea of cicada taken together with the low, but significant values from cornea and intersegmental membrane from adult locusts (Andersen, 1974b) indicate that these samples of soft cuticles, often considered to be unsclerotized, are sclerotized in the epicuticle and maybe also in the procuticular layer immediately beneath the epicuticle. An attempt has been made to obtain a measure of the relative degree of sclerotization by determining the amounts of radioactivity incorporated into the various cuticular regions after injection of a single dose of labeled dopamine or NADA (Andersen, 1974b). This can give a picture of how the various regions compete for the available sclerotization precursor at the time of injection, but it cannot measure the total sclerotization occurring in the regions. Soon after ecdysis labeled dopamine was injected into nymphal and adult S. gregaria, and good agreement was observed between the amounts of radioactivity incorporated into the various cuticular regions and the amounts of ketocatechols recovered after acid hydrolysis of corresponding regions from noninjected animals. This indicates that yield of ketocatechols can be a useful measure of sclerotization of locust cuticle. The only exception was that significantly less ketocatechol was obtained from the adult mandibles than expected considering their ability to incorporate radioactive dopamine. The regional pattern of incorporation of labeled NADA into locust cuticle was the same as that observed for labeled dopamine, and both patterns changed similarly during maturation of the animals (Andersen, 1974b). No convincing correlation was observed between the rate at which the various cuticular regions released tritium from b-labeled NADA in vitro and either the uptake of labeled dopamine and NADA in vivo or the yield of ketocatechols obtained by hydrolysis of these regions. These results suggest that it is not the amounts of sclerotizing enzymes which are the main determining factor for the degree of sclerotization, but the local availability of sclerotizing precursors. Similar attempts to determine the rate limiting factors for cuticular sclerotization have, to the best of our knowledge, not been performed on other insect species.

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4.4.6. Comparative Aspects 4.4.6.1. Cuticular Darkening

It is an old observation that cuticular sclerotization often is accompanied by a darkening of the cuticle, resulting in various shades of brown or black coloration. The black colors are presumably always due to deposition of melanins formed by oxidation of dopamine or DOPA (see Section 4.4.3.2.1), and melanin formation can apparently be blocked without interference with sclerotization. Melanins are not deposited in the cuticle of an albino mutant of the locust S. gregaria, but cuticular sclerotization appears not to be affected (Malek, 1957; Karlson and Schlossberger-Raecke, 1962). Further, injection of the phenoloxidase inhibitor phenylthiourea into larvae of Protophormia terraenovae prevents melanin deposition during puparium formation but does not affect hardening or the appearance of brown color (Dennell, 1958). The formation of brown colors appears to be more directly linked to cuticular sclerotization, and various suggestions have been put forward to explain why some cuticles become brown during sclerotization while other cuticles remain colorless. It has been suggested that a correlation exists between the use of NBAD as a sclerotization precursor and the intensity of brown color of the sclerotized cuticle (Brunet, 1980; Morgan et al., 1987; Hopkins and Kramer, 1992), and it has also been suggested that a darkbrown cuticular color indicates that the ring system of the sclerotization precursors is linked directly to cuticular proteins via quinone formation, whereas formation of links between the NADA side chain and the proteins results in a colorless cuticle (Andersen, 1974a). That suggestion was based upon the observation that colorless or lightly colored cuticles preferentially release tritium from the side chain of NADA and that dark-brown cuticles preferentially release tritium from the ring positions. The colorless benzodioxine-type compounds, which on acid hydrolysis yield ketocatechols, are preferentially formed from NADA and only to a minor extent from NBAD, as NBAD appears to be a poor substrate for the enzymatic activities responsible for introducing a double bond into the side chain (Andersen, 1989d). During sclerotization NBAD can be expected to form links to cuticular proteins via both ring positions and the b-position of the side chain, and some of the b-alanyl amino groups may react with quinones, while most of the NADA residues will be processed to dehydro-NADA, resulting in formation of colorless, protein-linked dihydroxyphenyl-benzodioxine derivatives. The formation of

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brown color during sclerotization can thus depend both on the amounts of NBAD available and the balance between quinone-forming enzymes and the enzyme activity catalyzing formation of a double bond in the side chain (Andersen, 1989b).

combination of individual reactions that have been characterized in different types of cuticles, and we have not yet obtained sufficient evidence that all these reactions, and only these reactions, are of general occurrence in cuticular sclerotization.

4.4.6.2. Cuticular Sclerotization in Insects Compared to That in Other Arthropods

4.4.7.1. Alternative Pathway for Dehydro-NADA Formation?

The release of ketocatechols during acid hydrolysis can be used as an indication of the involvement of dehydro-NADA in sclerotization, and makes it possible to determine how widespread the occurrence of this variant of cuticular sclerotization is. Ketocatechols have been obtained in varying amounts from cuticular samples of all pterygote insects studied so far, and especially the wings were found to be a good source of ketocatechols. None of the apterygote insects analyzed, representing Thysanura, Collembola, and Diplura, gave any measurable amounts of ketocatechols, and neither did the sclerotized cuticle of noninsectan arthropods, such as Decapoda, Isopoda, Araneae, Xiphosura, and Acarina (Andersen, 1985). The distribution of ketocatechol-yielding material among cuticles indicates that development of the ability to fly occurred in parallel with the development of the use of dehydro-NADA in sclerotization, resulting in a form of sclerotin which combines strength, toughness, and lightness to an optimal degree for flight purposes. The suggestion needs to be investigated in much more detail, and a detailed characterization of the sclerotization process(es) in cuticle of noninsectan arthropods is also needed. Little is known of sclerotization in crustaceans or most other arthropod groups.

In Figure 2 the sclerotization process is depicted as a linear reaction chain, where reactive intermediates formed at different steps in the chain can react with cuticular proteins, and it is supposed that the sclerotization patterns in different cuticular types depend partly upon the absolute and relative amounts of the two sclerotization precursors, NADA and NBAD, entering the chain at the beginning of the process and partly upon the extent to which the intermediate products will react with cuticular proteins or will continue along the reaction chain. If the o-quinone isomerase is present in larger amounts than the p-quinone methide isomerase one should expect that the quinone methide will be produced more rapidly than it is isomerized, favoring reaction between the quinone methide and proteins, whereas if the opposite is the case only a small fraction of the NADA p-quinone methide produced will have an opportunity to react with proteins before being isomerized to dehydro-NADA. Inhibition of laccase in locust cuticle by Na-azide results in a decrease in the consumption of NADA and an accumulation of dehydro-NADA in the incubation medium. As dehydro-NADA is a better substrate for the cuticular laccase than NADA, it was unexpected that Na-azide inhibits the laccasecatalyzed oxidation of dehydro-NADA much more than oxidation of NADA (Andersen, 1989a). One possible explanation could be that dehydro-NADA can be formed not only by isomerization of the p-quinone methide, but directly from NADA by means of a special enzyme, a desaturase, as suggested by Andersen et al. (1996). Another possible explanation could be that different enzymes catalyze the oxidation of NADA and dehydro-NADA, and that the dehydro-NADA oxidizing enzyme is much more sensitive to azide inhibition than the NADA oxidizing enzyme. This could result in accumulation of dehydro-NADA, but will demand a strict compartmentalization for the two enzyme activities, as the surplus of dehydro-NADA should not have access to the NADA-oxidizing laccase.

4.4.7. Unsolved Problems The model shown in Figures 1 and 2 for sclerotization of insect cuticle can account for most of the observations and experimental results which have been published during the years of cuticular studies. It appears likely that we have now obtained an understanding of the main features of the chemical processes occurring during sclerotization, but some observations are difficult to reconcile with the suggested scheme. This may be due to faulty observations or errors in interpretation, but could also represent variations of the scheme developed to serve specialized demands in some types of cuticle, or may represent essential, but unrecognized elements in the general sclerotization scheme. In any case, these observations deserve critical study before we can consider our understanding of the chemistry of sclerotization complete. A weakness in the suggested sclerotization scheme is that it is a

4.4.7.2. Extracuticular Synthesis of Catechol– Protein Conjugates for Sclerotization?

Another question to be studied in more detail is to what extent protein-bound catecholic derivatives

Cuticular Sclerotization and Tanning

are transferred from hemolymph to cuticle to participate in sclerotization. It has been reported that the epidermal cells can transfer proteins, arylphorins, from hemolymph to the cuticular compartment (Scheller et al., 1980; Schenkel et al., 1983; Ko¨ nig et al., 1986; Peter and Scheller, 1991), and apparently such transfer can also occur for proteins to which catechols are covalently bound (Koeppe and Mills, 1972; Koeppe and Gilbert, 1974; Bailey et al., 1999). If the protein-bound catechols are oxidized to quinones inside the cuticle during sclerotization, it is reasonable to assume that they will react with residues in the other proteins present in the cuticle and thus participate in sclerotin formation, but so far it is not known for certain how, where, and whether such protein–catechol conjugates are formed, and whether such conjugates after transfer to the cuticle will take part in sclerotization. Diffusion problems and steric hindrance may make it difficult for the catecholic residues to access the cuticular diphenoloxidases, but it is possible that the catechols are oxidized by encountering small, easily diffusable quinones formed by enzyme catalyzed oxidation of free catechols. 4.4.7.3. Importance of Cuticular Dehydration?

Fraenkel and Rudall (1940) reported a significant decrease in cuticular water content in connection with puparium sclerotization in blowflies, and it was later argued that formation of covalent crosslinks cannot fully explain the changes in mechanical properties occurring during sclerotization and that controlled dehydration of the cuticular matrix may be the most important factor in the stabilization of cuticle (Hillerton and Vincent, 1979; Vincent and Hillerton, 1979; Vincent, 1980). Dehydration may be caused by increased hydrophobicity of cuticular proteins due to reaction with the enzymatically formed quinones, by filling the initially water-filled interstices between proteins with polymerized catechols, and by water being actively transported out of the cuticle by some transport system residing in the epidermal apical cell membrane. All three mechanisms are probably involved in cuticular dehydration. Water transport coupled to active transport of ions appears to be the process which can be most precisely controlled and may be the most important dehydration mechanism. 4.4.7.4. Lipids and Sclerotization?

Lipids may play an essential role in cuticular sclerotization (Wigglesworth, 1985, 1988), a possibility that should be studied in more detail. The epicuticle consists mainly of proteins and lipids connected to

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each other to form a thin, extremely resistant, and inextractable layer. It is not known how the lipids and proteins are linked together, but the resistance towards hydrolytic degradation indicates that it is not only ester bonds which are involved, and that stable carbon–carbon bonds between lipid molecules and between lipids and proteins may play an important role. Semiquinones and other free radicals can easily be formed during enzyme catalyzed oxidation of catechols to quinones, and free radicals may react with unsaturated lipids resulting in stable lipophenolic complexes. Such reactions could be part of the stabilization of the epicuticle, and they could also contribute to making the connections between epicuticle and the underlying procuticle more stable and secure.

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4.5 Biochemistry of Digestion W R Terra and C Ferreira, University of Sa˜o Paulo, Sa˜o Paulo, Brazil ß 2005, Elsevier BV. All Rights Reserved.

4.5.1. Introduction 4.5.2. Overview of the Digestive Process 4.5.2.1. Initial Considerations 4.5.2.2. Characterization of Digestive Enzymes 4.5.2.3. Classification of Digestive Enzymes 4.5.2.4. Phases of Digestion and Their Compartmentalization in the Insect Gut 4.5.2.5. Role of Microorganisms in Digestion 4.5.3. Midgut Conditions Affecting Enzyme Activity 4.5.4. Digestion of Carbohydrates 4.5.4.1. Initial Considerations 4.5.4.2. Amylases 4.5.4.3. b-Glucanases 4.5.4.4. Xylanases and Pectinases 4.5.4.5. Chitinases and Lysozymes 4.5.4.6. a-Glucosidases 4.5.4.7. b-Glucosidases and b-Galactosidases 4.5.4.8. Trehalases 4.5.4.9. Acetylhexosaminidases, b-Fructosidases, and a-Galactosidases 4.5.5. Digestion of Proteins 4.5.5.1. Initial Considerations 4.5.5.2. Serine Proteinases 4.5.5.3. Cysteine Proteinases 4.5.5.4. Aspartic Proteinases 4.5.5.5. Aminopeptidases 4.5.5.6. Carboxypeptidases and Dipeptidases 4.5.6. Digestion of Lipids and Phosphates 4.5.6.1. Overview 4.5.6.2. Lipases 4.5.6.3. Phospholipases 4.5.6.4. Phosphatases 4.5.7. The Peritrophic Membrane 4.5.7.1. The Origin, Structure, and Formation of the Peritrophic Membrane 4.5.7.2. The Physiological Role of the Peritrophic Membrane 4.5.8. Organization of the Digestive Process 4.5.8.1. Evolutionary Trends of Insect Digestive Systems 4.5.8.2. Digestion in the Major Insect Orders 4.5.9. Digestive Enzyme Secretion Mechanisms and Control 4.5.10. Concluding Remarks

4.5.1. Introduction Most reviews start with the statement that the field under study has undergone remarkable progress over the last decade and the same can be said about the biochemistry of insect digestion. This growth is a characteristic of science as a whole that on average doubles in size every 15 years (Price, 1963). The growth of knowledge in the biochemistry of insect digestion had a bright start during the first decades of the last century, but slowed down

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after the development of synthetic chemical insecticides in the 1940s. Later on, with the environmental problems caused by chemical insecticides, new approaches for insect control were investigated. Midgut studies were particularly stimulated after the realization that the gut is a very large and relatively unprotected interface between the insect and its environment. Hence, an understanding of gut function was thought to be essential when developing methods of control that act through the gut,

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such as the use of transgenic plants to control phytophagous insects. Applebaum (1985) in his review on the biochemistry of digestion for the first edition of this series recognized the beginning of the renewed growth of the field. He discussed contemporary research showing that most insect digestive enzymes are similar to their mammalian counterparts, but that insect exotic diets require specific enzymes. In the next decade it became apparent that even enzymes similar to those of mammals have distinct characteristics because each insect taxon deals with food in a special way (Terra and Ferreira, 1994). Since then, the field of digestive physiology and biochemistry has progressed dramatically at the molecular level. The aim of this chapter is to review the recent and spectacular progress in the study of insect digestive biochemistry. To provide a broad coverage while keeping the chapter within reasonable size limits, only a brief account with key references is given for work done prior to 1994. Papers after 1994 have been selected from those richer in molecular details, and, when they were too numerous, representative papers were chosen, especially when rich in references to other papers. Throughout, the focus is on providing a coherent picture of phenomena and highlighting further research areas. Amino acid residues are denoted by the one-letter code, if in peptides, for the sake of brevity. When mentioned in text with a position number, amino acid residues are denoted by the three-letter code to avoid ambiguity. For consistency, traditional abbreviations, like BAPA for benzoyl-arginine p-nitroanilide, have been changed, in the example to B-R-pNA, because the one-letter code for arginine is R. The chapter is organized into four parts. The first part (Sections 4.5.2 and 4.5.3) tries to establish uniform parameters for studying insect digestive enzymes, providing an overview of the biochemistry of insect digestion, and discusses factors affecting digestive enzymes in vivo. The second part (Sections 4.5.4–4.5.6) reviews digestive enzymes with emphasis on molecular aspects, whereas the third part (Sections 4.5.7 and 4.5.8) describes the details of the digestive biochemical process along insect evolution. Finally, the fourth part (Section 4.5.9) discusses data on digestive enzyme secretion mechanisms and control.

4.5.2. Overview of the Digestive Process 4.5.2.1. Initial Considerations

Digestion is the process by which food molecules are broken down into smaller molecules that are absorbed by cells in the gut tissue. This process is

controlled by digestive enzymes and is dependent on their localization in the insect gut. 4.5.2.2. Characterization of Digestive Enzymes

Enzyme kinetic parameters are meaningless unless assays are performed in conditions in which enzymes are stable. If researchers adopt uniform parameters and methods, comparisons among similar and different insect species will be more meaningful. A rectilinear plot of product formation (or substrate disappearance) versus time will ensure that enzymes are stable in a given condition. Activities (velocities) calculated from this plot are reliable parameters. According to the International Union of Biochemistry and Molecular Biology, the assay temperature should be 30
C, except when the enzyme is unstable at this temperature or altered for specific purposes. Owing to partial inactivation, the optimum temperature is not a true property of enzymes and therefore should not be included in the characterization. Enzyme pH optimum should be determined using different buffers to discount the effects of chemical constituents of the buffers and their ionic strength on enzyme activity. The number of molecular forms of a given enzyme should be evaluated by submitting the enzyme preparation to a separation process (gel permeation, ion-exchange chromatography, electrophoresis, gradient ultracentrifugation, etc.), followed by assays of the resulting fractions. Substrate specificity of each molecular form of a given enzyme should be evaluated and substrate preference quantified by determining Vm/Km ratios for each substrate, keeping the amount of each enzyme form constant. Substrate preference expressed as the percentage activity towards a given substrate in relation to the activity upon a reference substrate may be misleading because, in this condition, enzyme activities are determined at different substrate saturations. The isoelectric points of many enzymes can be determined after staining with specific substrates following the separation of the native enzymes on isoelectrofocusing gels. If enzyme characterization is performed as part of a digestive physiology study, emphasis should be given to enzyme compartmentalization, substrate specificity, and substrate preference, in order to discover the sequential action of enzymes during the digestive process. Knowledge of the effect of pH on enzyme activity is useful in evaluating enzyme action in gut compartments (Figure 1) with different pH values. Finally, the determination of the molecular masses of digestive enzymes, associated with the ability of enzymes to pass through the peritrophic membrane, allows estimation of the pore sizes of the peritrophic membrane. Molecular

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Figure 1 Diagrammatic representation of insect gut compartments. Glycocalyx: the carbohydrate moiety of intrinsic proteins and glycolipids occurring in the luminal face of microvillar membranes.

masses determined in nondenaturing conditions are preferred, since in these conditions the enzymes should maintain their in vivo aggregation states (not only their quaternary structures if present). The method of choice in this case is gradient ultracentrifugation. Complete enzymological characterization requires purification to homogeneity and sequencing. Furthermore, details of the catalytic mechanisms, including involvement of amino acid residues in catalysis and substrate specificity, should be determined. This permits the classification of insect digestive enzymes into catalytic families, and discloses

the structural basis of substrate specificities; it will also enable us to establish evolutionary relationships with enzymes from other organisms. Cloning cDNA sequences encoding digestive enzymes enables the expression of large amounts of recombinant enzymes that may be crystallized or used for the production of antibodies. Antibodies are used in Western blots to identify a specific enzyme in protein mixtures or to localize the enzyme in tissue sections in a light or electron microscope. Enzyme crystals used for resolving threedimensional (3D) structures (via X-ray diffraction or nuclear magnetic resonance (NMR)) need amounts

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of purified enzymes that frequently are difficult to isolate from insects by conventional separation procedures. However, detailed 3D structures are necessary to understand enzyme mechanisms and the binding of inhibitors to enzyme molecules. Alternatively, cDNA may be amenable to site-directed mutagenesis for structure/function studies. Sitedirected mutagenesis tests the role of individual amino acid residues in enzyme function or structure. Such knowledge is a prerequisite in developing new biotechnological approaches to control insects via the gut. 4.5.2.3. Classification of Digestive Enzymes

Digestive enzymes are hydrolases. The enzyme classification and numbering system used here is that recommended by the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (Enzyme Commission). Peptidases (peptide hydrolases, EC 3.4) are enzymes that act on peptide bonds and include the proteinases (endopeptidases, EC 3.4.21–24) and the exopeptidases (EC 3.2.4.11–19). Proteinases are divided into subclasses on the basis of catalytic mechanism, as shown with specific reagents or effect of pH. Specificity is only used to identify individual enzymes within subclasses. Serine proteinases (EC 3.4.21) have a serine and a histidine in the active site. Cysteine proteinases (EC 3.4.22) possess a cysteine in the active site and are inhibited by mercurial compounds. Aspartic proteinases (EC 3.4.23) have a pH optimum below 5, due to the involvement of a carboxyl residue in catalysis. Metalloproteinases (EC 2.3.24) need a metal ion in the catalytic process. Exopeptidases include enzymes that hydrolyze single amino acids from the N-terminus (aminopeptidases, EC 3.4.11) or from the C-terminus (carboxypeptidases, EC 3.4.16–18) of the peptide chain and those enzymes specific for dipeptides (dipeptide hydrolases, EC 3.4.13) (Figure 2). Glycosidases (EC 3.2) are classified according to their substrate specificities. They include the enzymes that cleave internal bonds in polysaccharides and are usually named from their substrates, e.g., amylase, cellulase, pectinase, and chitinase. They also include the enzymes that hydrolyze oligosaccharides and disaccharides. Oligosaccharidases and disaccharidases are usually named based on the monosaccharide that gives its reducing group to the glycosidic bond and on the configuration (a or b) of this bond (Figure 2). Lipids are a large and heterogeneous group of substances that are relatively insoluble in water but readily soluble in apolar solvents. Some contain fatty acids (fats, phospholipids, glycolipids, and

waxes) and others lack them (terpenes, steroids, and carotenoids). Ester bonds are hydrolyzed in lipids containing fatty acids before they are absorbed. The enzymes that hydrolyze ester bonds comprise: (1) carboxylic ester hydrolases (EC 3.1.1), e.g., lipases, esterases, and phospholipases A and B; (2) phosphoric monoester hydrolases (EC 3.1.3), which are the phosphatases; and (3) phosphoric diester hydrolases (EC 3.1.4), including phospholipases C and D (Figure 2). 4.5.2.4. Phases of Digestion and Their Compartmentalization in the Insect Gut

Most food molecules to be digested are polymers such as proteins and starch and are digested sequentially in three phases. Primary digestion is the dispersion and reduction in molecular size of the polymers and results in oligomers. During intermediate digestion, these undergo a further reduction in molecular size to dimers; in final digestion, they become monomers. Digestion usually occurs under the action of digestive enzymes from the midgut, with little or no participation of salivary enzymes. Any description of the spatial organization of digestion in an insect must relate the midgut compartments (cell, ecto-, and endoperitrophic spaces) to each phase of digestion and, hence, to the corresponding enzymes. To accomplish this, enzyme determinations must be performed in each midgut luminal compartment and in the corresponding tissue. Techniques of sampling enzymes from midgut luminal compartments and for identifying microvillar enzymes, and enzymes trapped in cell glycocalyx have been reviewed elsewhere (Terra and Ferreira, 1994). Frequently, initial digestion occurs inside the peritrophic membrane (see Sections 4.5.7.1 and 4.5.7.2), intermediate digestion in the ectoperitrophic space, and final digestion at the surface of midgut cells by integral microvillar enzymes or by enzymes trapped into the glycocalyx (Figure 1). Exceptions to this rule, and the procedures for studying the organization of the digestive process, will be detailed below. 4.5.2.5. Role of Microorganisms in Digestion

Most insects harbor a substantial microbiota including bacteria, yeast, and protozoa. Microorganisms might be symbiotic or fortuitous contaminants from the external environment. They are found in the lumen, adhering to the peritrophic membrane, attached to the midgut surface, or within cells. Intracellular bacteria are usually found in special cells, the mycetocytes, which may be organized in groups, the

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Figure 2 Digestion of important nutrient classes. Arrows point to bonds cleaved by enzymes. (a) Protein digestion; R, different amino acid moieties; (b) starch digestion; (c) b-linked glucoside; (d) lipid digestion; PL, phospholipase; R, fatty acyl moieties. (Reprinted with permission from Terra, W.R., 2003. Digestion. In: Resh, V.H., Carde´, R.T. (Eds.), Encyclopedia of Insects. Academic Press, San Diego, CA, pp. 310–313; ß Elsevier.)

mycetomes. Microorganisms produce and secrete their own hydrolases and cell death will result in the release of enzymes into the intestinal milieu. Any consideration of the spectrum of hydrolase activity in the midgut must include the possibility that some of the activity may derive from microorganisms. Despite the fact that digestive enzymes of some insects are thought to be derived from the microbiota, there are relatively few studies that show an unambiguous contribution of microbial hydrolases. Best examples are found among wood- and humusfeeding insects like termites, tipulid fly larvae, and scarabid beetle larvae. Although these insects may have their own cellulases (see Section 4.5.4.3.1), only fungi and certain filamentous bacteria

developed a strategy for the chemical breakdown of lignin. Lignin is a phenolic polymer that forms an amorphous resin in which the polysaccharides of the secondary plant cell wall are embedded, thus becoming protected from enzymatic attack (Terra et al., 1996; Brune, 1998; Dillon and Dillon, 2004). Microorganisms play a limited role in digestion, but they may enable phytophagous insects to overcome biochemical barriers to herbivory (e.g., detoxifying flavonoids and alkaloids). They may also provide complex-B vitamins for blood-feeders and essential amino acids for phloem feeders, produce pheromone components, or withstand the colonization of the gut by nonindigenous species (including pathogens) (Dillon and Dillon, 2004).

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4.5.3. Midgut Conditions Affecting Enzyme Activity The pH of the contents of the midgut is one of the important internal environmental properties that affect digestive enzymes. Although midgut pH is hypothesized to result from adaptation of an ancestral insect to a particular diet, its descendants may diverge, feeding on different diets, while still retaining the ancestral midgut pH condition. Thus there is not necessarily a correlation between midgut pH and diet. In fact midgut pH correlates well with insect phylogeny (Terra and Ferreira, 1994; Clark, 1999). The pH of insect midgut contents is usually in the 6–7.5 range. Major exceptions are the very alkaline midgut contents (pH 9–12) of Lepidoptera, scarab beetles and nematoceran Diptera larvae, the very acid (pH 3.1–3.4) middle region of the midgut of cyclorrhaphous Diptera, and the acid posterior region of the midgut of heteropteran Hemiptera (Terra and Ferreira, 1994; Clark, 1999). pH values may not be equally buffered along the midgut. Thus, midgut contents are acidic in the anterior midgut and nearly neutral or alkaline in the posterior midgut in Dictyoptera, Orthoptera, and most families of Coleoptera. Cyclorrhaphan Diptera midguts have nearly neutral contents in the anterior and posterior regions, whereas in middle midgut the contents are very acid (Terra and Ferreira, 1994). The high alkanity of lepidopteran midgut contents is thought to allow these insects to feed on plant material rich in tannins, which bind to proteins at lower pH, reducing the efficiency of digestion (Berenbaum, 1980). This explanation may also hold for scarab beetles and for detritus-feeding nematoceran Diptera larvae that usually feed on refractory materials such as humus. Nevertheless, mechanisms other than high gut pH must account for the resistance to tannin displayed by some locusts (Bernays et al., 1981) and beetles (Fox and Macauley, 1977). One possibility is the effect of surfactants, such as lysolecithin which is formed in insect fluids due to the action of phospholipase A on cell membranes (Figure 2), and which occurs widely in insect digestive fluids (De Veau and Schultz, 1992). Surfactants are known to prevent the precipitation of proteins by tannins even at pH as low as 6.5 (Martin and Martin, 1984). Present knowledge is not sufficient to relate midgut detergency to diet or phylogeny or to both. Tannins may have deleterious effects other than precipitating proteins. Tannic acid is frequently oxidized in the midgut lumen, generating peroxides, including hydrogen peroxide, which readily diffuses

across cell membranes and is a powerful cytotoxin. In some insects, e.g., Orgyia leucostigma, tannic acid oxidation and the generation of peroxides are suppressed by the presence of high concentrations of ascorbate and glutathione in the midgut lumen (Barbehenn et al., 2003). Dihydroxy phenolics in an alkaline medium are converted to quinones that react with lysine e-amino groups. This leads to protein aggregation and a decrease in lysine availability for the insect. Other compounds, e.g., oleuropein, alkylate lysine residues in proteins, causing the same problems as dihydroxy phenolics. These phenomena are inhibited in larvae of several lepidopteran species by secreting glycine into the midgut lumen. Glycine competes with lysine residues in the denaturating reaction (Konno et al., 2001). A high midgut pH may also be of importance, in addition to its role in preventing tannin binding to proteins, in freeing hemicelluloses from plant cell walls ingested by insects. Hemicelluloses are usually extracted in alkaline solutions for analytical purposes (Blake et al., 1971) and insects, such as the caterpillar Erinnyis ello, are able to digest hemicelluloses efficiently without affecting the cellulose from the leaves they ingest to any degree (Terra, 1988). This explanation is better than the previous one in accounting for the very high pH observed in several insects, since a pH of about 8 is sufficient to prevent tannin binding to proteins (Terra, 1988). The acid region in the cyclorrhaphous Diptera midgut is assumed to be involved in the process of killing and digesting bacteria, which may be an important food for maggots. This region is retained in Muscidae that have not diverged from the putative ancestral bacteria-feeding habit, as well as in the flesh-feeding Calliphoridae and in the fruit-feeding Tephritidae (Terra and Ferreira, 1994). The acid posterior midgut of Hemiptera may be related to their lysosome-like digestive enzymes (cysteine and aspartic proteinase) (see Sections 4.5.5.3 and 4.5.5.4). Few papers have dealt with midgut pH buffering mechanisms. The early unsuccessful attempts to relate midgut buffering activity to the large amounts of phosphate frequently found in insect midguts, as well as other unsuccessful attempts to describe buffering mechanisms, are reviewed by House (1974). The results of more recent research on midgut buffering mechanisms are more encouraging. Dow (1992) showed that the lepidopteran larval midgut transports equal amounts of Kþ and alkali from blood to the midgut lumen. Based on this and other data he described a carbonate secretion system, which may be responsible for the high pH found in Lepidoptera midguts (Figure 3). Phosphorus NMR

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Figure 3 A model for generation of high gut pH by the goblet cells of lepidopteran larvae. Carbonic anhydrase (CA) produces carbonic acid that dissociates into bicarbonate and a proton. The proton is pumped by a V-ATPase into the goblet cell cavity, from where it is removed in exchange with Kþ that eventually diffuses into lumen. Bicarbonate is secreted in exchange with chloride and loses a proton due to the intense field near the membrane, forming carbonate and raising the gut pH. (Data from Dow, J.A.T., 1992. pH gradients in lepidopteran midgut. J. Exp. Biol. 172, 355–375.)

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microscopy has been used to show that valinomycin leads to a loss of alkalinization in the midgut of Spodoptera litura (Skibbe et al., 1996). As valinomycin is known to transport Kþ down its concentration gradient, this result gives further support to the model described in Figure 3. It is not known whether midgut alkalinization in scarab beetles and nematoceran Diptera occurs by mechanisms similar to those of lepitopteran larvae. Terra and Regel (1995) determined pH values and concentrations of ammonia, chloride, and phosphate in the presence or absence of ouabain and vanadate in Musca domestica midguts. From the results they proposed that middle midgut acidification is accomplished by a proton pump of mammalian-like oxyntic cells, whereas the neutralization of posterior midgut contents depends on ammonia secretion (Figure 4). Redox conditions in the midgut are regulated and may be the result of phylogeny, although data are scarce. Reducing conditions are observed in clothes moth, sphinx moths, owlet moths, and dermestid beetles (Appel and Martin, 1990) and in Hemiptera (Silva and Terra, 1994). Reducing conditions are important to open disulfide bonds in keratin ingested by some insects (clothes moths, dermestid beetles) (Appel and Martin, 1990), to maintain the activity of the major proteinase in Hemiptera (see Section 4.5.5.3), and to reduce the impact of some plant allelochemicals, such as phenol, in some herbivores (Appel and Martin, 1990). In spite of this, the artificial lowering of in vivo redox potentials did not

Figure 4 Diagrammatic representation of ion movements, proposed as being responsible for maintenance of pH in the larval midgut contents of Musca domestica. Carbonic anhydrase (CA) in cup-shaped oxyntic cells in the middle of the midgut (a) produces carbonic acid which dissociates into bicarbonate and a proton. Bicarbonate is transported into the hemolymph, whereas the proton is actively translocated into the midgut lumen acidifying its contents to pH 3.2. Chloride ions follow the movement of protons. NH3 diffuses from anterior and posterior midgut cells (b) into the midgut lumen, becoming protonated and neutralizing their contents to þ þ þ þ pH 6.1–6.8. NHþ 4 is then exchanged for Na by a microvillar Na /K -ATPase. Inside the cells, NH4 forms NH3, which diffuses into midgut lumen, and proton that is transferred into the hemolymph. (Reprinted with permission from Terra, W.R., Regel, R., 1995. pH buffering in Musca domestica midguts. Comp. Biochem. Physiol. A 112, 559–564; ß Elsevier.)

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significantly impact digestive efficiency of the herbivore Helicoverpa zea, although the reducing agent used (dithiothreitol) inhibited some proteinases in vitro (Johnson and Felton, 2000). Although several allelochemicals other than phenols may be present in the insect gut lumen, including alkaloids, terpene aldehydes, saponins, and hydroxamic acids (Appel, 1994), there are no sufficient data on their effect on digestion.

4.5.4. Digestion of Carbohydrates 4.5.4.1. Initial Considerations

Polysaccharides are major constituents of cell walls and energy reserves such as starch granules within plant cells and glycogen within animal cells. For phytophagous insects, disruption of plant cell walls is necessary in order to expose storage polymers in cell contents to polymer hydrolases. Cell wall breakdown may be achieved by mastication, but more frequently it is the result of the action of digestive enzymes. Thus, even insects unable to obtain nourishment from the cellulosic and noncellulosic cell wall biochemicals would profit from having enzymes active against these structural components. Cell walls are disrupted by b-glucanases, xylanases, and pectinases (plant cells), lysozyme (bacterial cells), or chitinase and b-1,3-glucanase (fungal cells). The carbohydrates associated with cellulose are frequently named ‘‘hemicelluloses’’ and the enzymes that attack them ‘‘hemicellulases.’’ Thus, the hemicellulases include b-glucanases other than cellulases, xylanases, and pectinases. Following the loss of cell wall integrity, starch digestion is initiated by amylases. A complex of carbohydrases converts the oligomers resulting from the action of the polymer hydrolyzing enzymes into dimers (such as sucrose, cellobiose, and maltose that also occur free in some cells) and finally into monosaccharides like glucose and fructose. 4.5.4.2. Amylases

a-Amylases (EC 3.2.1.1) catalyze the endohydrolysis of long a-1,4-glucan chains such as starch and glycogen. Amylases are usually purified by glycogen–amylase complex formation followed by precipitation in cold ethanol or, alternatively, by affinity chromatography in a gel matrix linked to a protein amylase inhibitor. In sequence, isoamylases can be resolved by anion exchange chromatography (Terra and Ferreira, 1994). Most insect amylases have molecular weights in the range 48–60 kDa, pI values of 3.5–4.0, and Km values with soluble starch around 0.1%. pH optima generally correspond to the pH prevailing in midguts from which the amylases were isolated. Insect

amylases are calcium-dependent enzymes and are activated by chloride with displacement of the pH optimum. Activation also occurs with anions other than chloride, such as bromide and nitrate, and it seems to depend upon the ionic size (Terra and Ferreira, 1994). The best-known insect a-amylase, and the only one whose 3D structure has been resolved, is the midgut a-amylase of Tenebrio molitor larvae. The enzyme has three domains. The central domain (domain A) is an (b/a)8-barrel that comprises the core of the molecule and includes the catalytic amino acid residues (Asp 185, Glu 222, and Asp 287) (T. molitor a-amylase numbering throughout). Domains B and C are almost opposite to each other, on each side of domain A. The substrate-binding site is located in a long V-shaped cleft between domains A and B. There, six saccharide units can be accommodated, with the sugar chain being cleaved between the third and fourth bound glucose residues. A calcium ion is placed in domain B and is coordinated by Asn 98, Arg 146, and Asp 155. This ion is important for the structural integrity of the enzyme and seems also to be relevant due its contact with His 189. This residue interacts with the fourth sugar of the substrate bound in the active site, forming a hinge between the catalytic site and the Ca2þbinding site. A chloride ion is coordinated by Arg 183, Asn 285, and Arg 321 in domain A (Strobl et al., 1998a). Domain C is placed in the C-terminal part of the enzyme, contains the so-called ‘‘Greek key’’ motif and has no clear function (Figure 5). These structural features are shared by all known a-amylases (Nielsen and Borchert, 2000), although not all a-amylases have a chloride-binding site (Strobl et al., 1998a). The most striking difference between mammalian and insect a-amylases is the presence of additional loops in the vicinity of the active site of the mammalian enzymes (Strobl et al., 1998a). Asp 287 is conserved in all a-amylases. Comparative studies have shown that Glu 222 is the proton donor and Asp 185 the nucleophile, and that Asp 287 is important but not a direct participant in catalysis. It is proposed that its role is to elevate the pKa of the proton donor (Nielsen and Borchert, 2000). Chloride ion is an allosteric activator of a-amylases, leading to a conformation change in the enzyme that changes the environment of the proton donor. This change causes an increase in the pKa of the proton donor, thus displacing the pH optimum of the enzyme and increasing its Vmax (Levitzki and Steer, 1974). According to Strobl et al. (1998a), the increase in Vmax is a consequence of chloride ion being close to the water molecule that has been

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Figure 5 Conserved residues in the primary structures of major insect digestive enzymes. AMY (amylase) follows Tenebrio molitor amylase numbering; CHI (chitinase), molting-fluid Manduca sexta chitinase numbering. TRY (trypsin) and CHY (chymotrypsin) follow the bovine chymotrypsin numbering; CAL (cathepsin L), papain numbering: APN (aminopeptidase N) does not have a consensual numbering; CPA (carboxypeptidase A), mammalian CPA numbering.

suggested to initiate the cleavage of the substrate chain. The nucleophilicity of this water molecule might be enhanced by the negative charge of the ion. There are 21 complete sequences of insect a-amylases registered in the GenBank (as of May 2003) excluding all of the sequences of Drosophila. The sequences correspond to 18 species found in four orders. Examples may be found among Hymenoptera (Ohashi et al., 1999; Da Lage et al., 2002), Coleoptera (Strobl et al., 1997; Titarenko and Chrispeels, 2000), Diptera (Grossmann et al., 1997; Charlab et al., 1999), and Lepidoptera (Da Lage et al., 2002). All the sequences have the catalytic triad (Asp 185, Glu 222, and Asp 287), the substrate-binding histidine residues (His 99, His 189, and His 286), and the Ca2þ-coordinating residues (Asn 98, Arg 146, and Asp 155) (Figure 5). From the residues found to be involved in chloride binding, Arg 183 and Asn 285 are conserved, whereas

position 321 varies. According to D’Amico et al. (2000), all known chloride-activated a-amylases have an arginine or lysine residue at position 321. Insect a-amylase sequences have arginine at position 321, except those of Zabrotes subfasciatus and Anthonomus grandis, which have lysine, and the lepidopteran a-amylases which have glutamine. This agrees with the observation that most insect a-amylases are activated by chloride with the remarkable exception of lepidopteran amylases (Terra and Ferreira, 1994). The few coleopteran and hymenopteran a-amylases reported to be not affected by chloride (Terra and Ferreira, 1994) deserve reinvestigation. It is possible that another anion is replacing chloride as an activator, as shown for some hemipteran amylases (Hori, 1972). Action pattern refers to the number of bonds hydrolyzed during the lifetime of a particular enzyme–substrate complex. If more than one bond

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is hydrolyzed after the first hydrolytic step, the action pattern is said to be processive. The degree of multiple attack is the average number of hydrolyzed bonds after the first bond is broken. Rhynchosciara americana amylase has a degree of multiple attack between that of the amylase of Bacillus subtilis (1.7) and porcine pancreas (6). Amylases from larvae and adults of Sitophilus zeamais, S. granarius, and S. oryzae and larvae of Bombyx mori have action patterns similar to that of porcine pancreas amylase (Terra and Ferreira, 1994). These studies need to be taken further, including the determination of the affinities corresponding to each subsite in the active center. Such studies, specially if combined with crystallographic data, may describe in molecular detail the reasons why amylases act differently toward starches of distinct origins. There is a variety of natural compounds that affect amylases, including many plant protein inhibitors (Franco et al., 2002). Crystallographic data have shown that these protein inhibitors always occupy the amylase active site (Strobl et al., 1998b; Nahoum et al., 1999). In the case of the Amaranth a-amylase inhibitor, a comparison of T. molitor amylase–inhibitor complex with a modeled complex between porcine pancreatic a-amylase and the inhibitor identified six hydrogen bonds that can be formed only in the T. molitor amylase–inhibitor complex (Pereira et al., 1999). This was the first successful explanation of how a protein inhibitor specifically inhibits a-amylases from insects, but not from mammalian sources. As will be discussed with details for trypsins (see Section 4.5.5.2.1), specific amylases are induced when insect larvae are fed with a-amylase inhibitor-containing diets (Silva et al., 2001). The mechanisms underlying this induction are unknown and are presently under study (C.P. Silva, personal communication). 4.5.4.3. b-Glucanases

b-Glucanases are enzymes acting on b-glucans. These are major polysaccharide components of plant cell walls and include b-1,4-glucans (cellulose), b-1,3-glucans (callose), and b-1,3;1,4-glucans (cereal b-glucans) (Bacic et al., 1988). The cell walls of certain groups of fungi have b-1,3;1,6-glucans (Bacic et al., 1988). 4.5.4.3.1. Cellulases Cellulose is by far the most important b-glucan. It occurs in the form of b-1,4glucan chains packed in an ordered manner to form compact aggregates which are stabilized by hydrogen bonds. The resulting structure is crystalline and not soluble. According to work done with microbial systems, cellulose is digested by a

combined action of two enzymes. An endo-b-1,4glucanase (EC 3.2.1.4), with an open substratebinding cleft, cleaves bonds located within chains in the amorphous regions of cellulose, creating new chain ends. An exo-b-1,4-glucanase (EC 3.2.1.91) processively releases cellobiose from the ends of cellulose chains in a tunnel-like active site. Surface loops in cellobiohydrolase prevent the dislodged cellulose chains from readhering to the crystal surface, as the enzyme progresses into crystalline cellulose (Rouvinen et al., 1990; Kleywegt et al., 1997). Cellobiohydrolase structure is modular, comprising a catalytic domain linked to a distinct cellulosebinding domain, which enhances the activity of the enzyme towards insoluble cellulose (Linder and Teeri, 1997). Cellulose digestion in insects is rare, presumably because the dietary factor that usually limits growth in plant feeders is protein quality. Thus, cellulose digestion is unlikely to be advantageous to an insect that can meet its dietary requirements using more easily digested constituents. Nevertheless, cellulose digestion occurs in several insects that have, as a rule, nutritionally poor diets (Terra and Ferreira, 1994). The role of symbiotic organisms in insect cellulose digestion is less important than initially believed (Slaytor, 1992), although symbiotic nitrogen-fixing organisms are certainly involved in increasing the nutritive value of diets of many insects (Terra, 1990). Few insect cellulases have been purified and characterized. Two endo-b-1,4-glucanases (41 and 42 kDa) were isolated from the lower termite Reticulitermes speratus (Watanabe et al., 1997). The cDNA that encodes this protein was cloned (Watanabe et al., 1998) and the protein was shown to be secreted from the salivary glands (Tokuda et al., 1999). The cDNAs that encode the endo-b1,4-glucanase secreted by the midgut of the higher termite Nasutitermes takasagoensis (Tokuda et al., 1999) and the two endo-b-1,4-glucanases from the woodroach Panestria cribrata of 47 kDa (Tokuda et al., 1997) and 54 and 49 kDa (Scrivener and Slaytor, 1994), respectively, were also cloned (Tokuda et al., 1999; Lo et al., 2000). Alignments of the sequences of termite and woodroach endoglucanases from data banks showed that they belong to family 9 of glycoside hydrolases (Coutinho and Henrissat, 1999). The paradigm of this family is the endo/exocellulase from the bacteria Thermomonospora fusca, whose catalytic center binds a cellopentaose residue and cleaves it into cellotetraose plus glucose or cellotriose plus cellobiose, and has Asp 55 as a nucleophile and Glu 424 as a proton donor (Sakon et al., 1997).

Biochemistry of Digestion

The active site groups are conserved in the termite and woodroach endoglucanases although these enzymes lack the cellulose-binding domains that improve the binding and facilitate the activity of the catalytic domain on crystalline cellulose (Linder and Teeri, 1997). The conclusions drawn from sequence alignments were confirmed by the 3D structure resolution of the N. takasagoensis endoglucanase, which also revealed a Ca2þ-binding site near its substrate binding cleft (Khadeni et al., 2002). According to Slaytor (1992), the large production of endoglucanases in termites and woodroaches would compensate their low efficiency on crystalline cellulose. The phytophagous beetle Phaedon cochleariae has cellulase activity in its midgut (Girard and Jouanin, 1999a). A cDNA that encodes one cellulase was cloned and shown to belong to the glycoside hydrolase family 45 and to consist only of a catalytic domain. The purification of an exo-b-1,4glucanase (52 kDa) from the termite Macrotermes mulleri (Rouland et al., 1988) and the partial resolution from Ergates faber (Coleoptera) midgut extracts of another exo-b-1,4-glucanase (Chararas et al., 1983) suggest that these insects, in contrast to the others, have cellulases with cellulose-binding domains. 4.5.4.3.2. Laminarinases and licheninases Licheninases (EC 3.2.1.73) digest only b-1,3;1,4-glucans whereas laminarinases may hydrolyze b-1,3;1,4glucans and also b-1,3-glucans (EC 3.2.1.6) or only the last polymer (EC 3.2.1.39). In spite of laminarinase activities being widespread among insects (Terra and Ferreira, 1994), only the laminarinases of Periplaneta americana (LAM, LIC 1, and LIC 2) have been purified and studied in detail (Genta et al., 2003). The enzymes are secreted by salivary glands, stabilized by calcium ions, and have pH optima around 6. LAM (46.2 kDa) is an endo-b-1,3-glucanase that processively releases mainly glucose from laminarin and shows lytic activity on fungal cells. LIC 1 (24.6 kDa) is an endo-b-1,3-(4)-glucanase highly active on sequences of b-1,3-linked cellotetraoses, releasing cellotetraose from lichenin and is also able to lyse fungal cells. The specificities of P. americana b-glucanases agree with the omnivorous detritus-feeding habit of this insect, as they are able to help the cellulases in attacking plant cell walls (mainly those from cereals) and also in opening the cell walls of fungi, usually present in detritus. No insect licheninase has been characterized in detail to date.

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4.5.4.4. Xylanases and Pectinases

Xylans constitute the major noncellulosic polysaccharides (hemicelluloses) of primary walls of grasses and secondary walls of all angiosperms, accounting for one-third of all renewable organic carbon available on earth (Bacic et al., 1988). Chemically, xylans are a family of linear b-1,4-xylans with a few branches. Endo-b-1,4-xylanase activities (EC 3.2.1.8) were found in several insects (Terra and Ferreira, 1994). One of these enzymes was cloned from a beetle and shown to correspond to a protein of 22 kDa, with high sequence identity to fungal xylanases, and conserving the usual two catalytic regions (Girard and Jouanin, 1999a). An exo-b1,4-xylanase (EC 3.2.1.37) was partially purified from termites (Matoub and Rouland, 1995) and thought to act synergistically with an endo-b-1,4xylanase originating from fungus ingested by the termites. Much more work is needed on this class of enzymes that may be important mainly for detritivorous insects. Pectin is a linear chain of a d-galacturonic acid units with a-1,4-linkages in which varying proportions of the acid groups are present as methyl esters. It is the main component of the rhamnogalacturonan backbone of the structure formed by the pectin polysaccharides. Pectin is hydrolyzed by pectinases (polygalacturonases, EC 3.2.1.15) described in many insects (Terra and Ferreira, 1994). Pectinases are thought to be important for hemipterans, as they would facilitate penetration of their stylets through plant tissues into sap-conducting structures and for insects boring plant parts. Accordingly, pectinases have been found in hemipteran saliva (Vonk and Western, 1984) and have been isolated and characterized from two weevils (Shen et al., 1996; Dootsdar et al., 1997) and cloned from a phytophagous beetle (Girard and Jouanin, 1999a). The pectinases from the weevils S. oryzae (Shen et al., 1996) and Diaprepes abbreviatus (Dootsdar et al., 1997) were purified to electrophoretical homogeneity from whole-body extracts and gut homogenates, respectively. Purification of the pectinases was achieved by affinity chromatography through cross-linked pectate in addition to two ion exchange chromatographic steps. The enzymes have molecular masses of 35–45 kDa, pH optimum 5.5 and Km values of 1–4 mg ml1 for pectic acid. The D. abbreviatus pectinase is inhibited by a polygalacturonase-inhibitor protein that may be associated with plant resistance to insects (Dootsdar et al., 1997). Although the weevil pectinases may originate from endosymbiotic bacteria (Campbell et al., 1992), the finding that the cDNA-coding pectinase

182 Biochemistry of Digestion

of the beetle P. cochleariae has a poly(A) tail (Girard and Jouanin, 1999a) argues against this hypothesis. The beetle pectinase belongs to family 28 of the glycoside hydrolases and is most related to fungal endopolygalacturonases, conserving the active site signature centered on the His 223 catalytic residue (Girard and Jouanin, 1999a; Markovic and Janecek, 2001). 4.5.4.5. Chitinases and Lysozymes

Chitin, the simplest of the glycosaminoglycans, is a b-1,4-homopolymer of N-acetylglucosamine (see Chapter 4.3). Chitinolytic enzymes (Kramer and Koga, 1986) include: chitinase (EC 3.2.1.14), which catalyzes the random hydrolysis of internal bonds in chitin forming smaller oligosaccharides, and b-N-acetyl-d-glucosaminidase (EC 3.2.1.52), which liberates N-acetylglucosamine from the nonreducing end of oligosaccharides (Kramer and Koga, 1986). Lysozyme, as described below, also has some chitinase activity, whereas chitinase has no lysozyme activity. Chitinolytic enzymes associated with the ecdysial cycle have been studied and demonstrated to act synergistically in cuticular chitin degradation (Kramer and Koga, 1986). Nevertheless, these enzymes may also have a digestive role. Chitinase assays with midguts of several insects showed that there is a correlation between the presence of chitinase and a diet rich in chitin (Terra and Ferreira, 1994). The best-known insect chitinase is the molting fluid chitinase from the lepidopteran Manduca sexta (Figure 5). The enzyme has a multidomain architecture that includes a signal peptide, an N-terminal catalytic domain, with the consensus sequence (F/L)DG(L/I)DLDWEYP, and a C-terminal cysteine-rich chitin-binding domain, which are connected by the interdomain serine/threonine-rich O-glycosylated linker. The residues Asp 142, Asp 144, Trp 145, and Glu 146 of the consensus sequence have been shown by site-directed mutagenesis to be involved in catalysis. Glu 146 functions as a proton donor in the hydrolysis like homologous residues in other glycoside hydrolases. Asp 144 apparently functions as an electrostatic stabilizer of the positively charged transition state, whereas Asp 142, and perhaps also Trp 145, influences the pKa values of Asp 144 and Glu 146. The chitin-binding domains have 6 cysteines (with the consensus sequence CXnCXnCXnCXnCXnC, where Xn stands for a variable number of residues) and include several highly conserved aromatic residues (Tellam et al., 1999). The three disulfide bonds in the domain may constrain the polypeptide to present the aromatic amino acids on the protein surface for

interactions with the ring structures of sugars. Thus, the chitin-binding domains enhance activity toward the insoluble polymer, whereas the linker region facilitates secretion from the cell and helps to stabilize the enzyme in the presence of proteolytic enzymes (Kramer et al., 1993; Lu et al., 2002; Zhang et al., 2002; Arakane et al., 2003). Chitinase in molting fluid of the beetle T. molitor is a large (about 320 kDa) multidomain protein with five catalytic domains, five serine/threonine-rich linker domains, four chitin-binding domains, and two mucin-like domains. There is evidence that the enzyme is secreted as a zymogen activated by trypsin (Royer et al., 2002). The Anopheles gambiae gut chitinase is secreted upon blood-feeding as an inactive proenzyme that is later activated by trypsin. Sequencing a cDNA coding the gut chitinase showed that the enzyme comprises a putative catalytic domain at the N-terminus, a putative chitin-binding domain at the C-terminus, and a serine/threonine/proline-rich amino acid stretch between them (Shen and Jacobs-Lorena, 1997). The mosquito chitinase seems to modulate the thickness and permeability of the chitin-containing peritrophic membrane (see Section 4.5.7.1). Supporting this conjecture the authors found that the peritrophic membrane is stronger and persisted longer when the mosquitoes were fed diets containing chitinase inhibitor. The beetle P. cochleariae has one group of chitinases of 40–70 kDa active at pH 5.0 and detected in guts and another group of 40–70 kDa that are more active at pH 7.0 and that are associated with molting. A cDNA encoding a gut chitinase showed this enzyme has an active site centered on the catalytic residues Asp 146 and Glu 150 (M. sexta chitinase numbering), but lacks the C-terminal chitin-binding domain and the serine/threonine-rich interdomain (Girard and Jouanin, 1999b). The T. molitor midgut chitinase (GenBank accession no. AY325895) is similar to that of P. cochleariae. The putative role of P. cochleariae chitinase is in turnover of the peritrophic membrane (Girard and Jouanin, 1999b), as proposed above for A. gambiae. It is not clear, however, why the chitinase from A. gambiae in contrast to that of P. cochleariae has a chitin-binding domain if their putative role is to affect the same type (type I) (see Section 4.5.7.1) of peritrophic membrane. The T. molitor midgut chitinase may have the same proposed function, although it may also digest the cell walls of fungi usually present in its food. More research is necessary to clarify this point. Lysozyme (EC 3.2.1.17) catalyzes the hydrolysis of the 1,4-b-glycosidic linkage between N-acetylmuramic acid and N-acetylglucosamine of the

Biochemistry of Digestion

peptidoglycan present in the cell wall of many bacteria, causing cell lysis. Lysozyme is part of an immune defense mechanism against bacteria and has been described in most animals, including insects (Hultmark, 1996). Lysozyme has also been implicated in the midgut digestion of bacteria by organisms which ingest large amounts of them, such as marine bivalves (McHenery et al., 1979), or that harbor a bacterial culture in their guts, as exemplified by ruminants (Stewart et al., 1987). Among insects, the capacity of digesting bacteria in the midgut seems to be an ancestral trait of Diptera Cyclorrhapha (Lemos and Terra, 1991a; Regel et al., 1998), which agrees with the fact that most Diptera Cyclorrhapha larvae are saprophagous, feeding largely on bacteria (Terra, 1990). These insects have midgut lysozymes (Lemos et al., 1993; Regel et al., 1998) similar to those of vertebrate fermenters. Thus, these enzymes have low pI values, are more active at pH values 3–4, when present in media with physiological ionic strengths, and are resistant to the cathepsin D-like aspartic proteinase present in midguts (Lemos et al., 1993; Regel et al., 1998). Sequence analyses (Kylsten et al., 1992; Daffre et al., 1994; Ito et al., 1995) showed that cyclorrhaphan (Drosophila melanogaster and Musca domestica) digestive lysozymes have the same conserved residues as vertebrate lysozymes (Imoto et al., 1972) (numbering according to Regel et al., 1998): positions 55–61, Glu 36, and Asp 54. Glu 36 is believed to act as a general acid in catalysis, whereas Asp 54 is postulated to stabilize the resulting metastable oxocarbonium intermediate (Imoto et al., 1972). More recently, Asp 54 has been implicated more strongly in catalysis of the hydrolysis of chitinderived substrates (Matsumura and Kirsch, 1996). The ability of D. melanogaster and M. domestica purified lysozymes in hydrolyzing chitosan favors this view. The most remarkable sequence convergence of cyclorrhaphan digestive lysozymes with that of vertebrate foregut fermenters are Ser 104 and a decrease in the number of basic amino acids, suggesting that these features are necessary for a digestive role in an acid environment (Regel et al., 1998). These hypotheses must be tested by sitedirected mutagenesis and by the resolution of the 3D structure of at least one of these insect digestive lysozymes. Lysozyme is also found in the salivary glands of R. speratus. This insect is a termite that feeds mainly on dead wood, which tends to lack nitrogen. Fujita et al. (2001) suggested, on the basis of the distribution and activity of lysozyme in this termite, that wood-feeding termites use lysozyme secreted from

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the salivary gland to digest their hindgut bacteria, which are transferred by proctodeal trophallaxis. The termite lysozyme is active in neutral pH and has no serine in position 104 (Fujita et al., 2002), thus differing from the digestive cyclorrhaphan lysozymes. Chitin and the bacterial peptidoglycan resemble one another chemically and structurally. Because of this one might anticipate that chitinases and lysozymes would be structurally related enzymes. Indeed, some lysozymes, exemplified by cyclorrhaphan digestive lysozymes (Regel et al., 1998) are very good chitinases. Structurally well-known chitinases and lysozymes share no significant amino acid sequence similarities, but have a structurally invariant core consisting of two helices and a three-stranded b-sheet, which form the substratebinding catalytic cleft. These enzymes are considered to represent a superfamily of hydrolases which are likely to have arisen by divergent evolution (Monzingo et al., 1996). However, this picture does not take into account insect enzymes, for which 3D structural data are lacking. 4.5.4.6. a-Glucosidases

a-Glucosidases (EC 3.2.1.20) catalyze the hydrolysis of terminal, nonreducing a-1,4-linked glucose residues from aryl (or alkyl)-glucosides, disaccharides, or oligosaccharides. a-glucosidases are frequently named maltases, although some of them may have weak activity on maltose. Insect a-glucosidases occur as soluble forms in the midgut lumen or are trapped in the midgut cell glycocalyx. They are also bound to microvillar membranes (Terra and Ferreira, 1994), to perimicrovillar membranes (lipoprotein membranes ensheathing the midgut cell microvillar membranes in most hemipterans) (Silva and Terra, 1995), or to the modified perimicrovillar membranes of aphid midgut cells (Cristofoletti et al., 2003). The last two membrane-bound a-glucosidases, as well as the soluble enzyme from bee midguts (Nishimoto et al., 2001), were purified to electrophoretic homogeneity. Culex pipiens microvillar a-glucosidase is the primary target of the binary toxin of Bacillus sphericus and, although not purified, cDNA sequencing data suggest it is bound by a glycosyl phosphatidyl inositol anchor (Darboux et al., 2001). a-Glucosidase is a major protein in dipteran midgut microvillae (Terra and Ferreira, 1994) and probably because of that it is the receptor of endotoxins, similar to that observed with aminopeptidase N in lepidopteran midgut cells (see Section 4.5.5.5). Although biochemical properties of many crude, partially or completely purified gut a-glucosidases

184 Biochemistry of Digestion

are known, including molecular masses (range 60–80 kDa or a multiple of these values), pH optima (range 5–6.5, irrespective of the corresponding midgut pH value), pI values (range 5.0–7.2), and inhibition by tris(hydroxylmethyl)aminomethane (Tris), only few studies report on a-glucosidases specificities. These studies showed that insect a-glucosidases hydrolyze oligosaccharides up to at least maltopentaose (Terra and Ferreira, 1994), although there are exceptions. The perimicrovillar a-glucosidase from Dysdercus peruvianus prefers oligosaccharides up to maltotetraose (Silva and Terra, 1995) and the midgut bee a-glucosidase, up to maltotriose (Nishimoto et al., 2001). The purified midgut a-glucosidase of the pea aphid, Acyrthosiphon pisum catalyzes transglycosylation reactions in the presence of excess sucrose, thus freeing glucose from sucrose without increasing the osmolarity of the medium (Cristofoletti et al., 2003). This phenomenon associated with a quick fructose absorption (Ashford et al., 2000) explains why the midgut luminal osmolarity decreases as the ingested sucrose-containing phloem sap passes along the aphid midgut. Three digestive a-glucosidases from dipterans have been sequenced: one salivary (James et al., 1989) and two midgut (Zheng et al., 1995; Darboux et al., 2001) enzymes. All the sequences have the invariant residues: Asp 123, His 128, Asp 206, Arg 221, Glu 271, His 296, and Asp 297 (numbers are relative to the positions in the sequence of C. pipiens a-glucosidases) (Darboux et al., 2001) that are involved in the active site of a-amylase family of enzymes, and the three residues Gly 69, Pro 77, and Gly 323, that have a structural role for some a-glucosidases (Janecek, 1997). 4.5.4.7. b-Glucosidases and b-Galactosidases

b-Glycosidases (EC 3.2.1) catalyze the hydrolysis of terminal, nonreducing b-linked monosaccharide residues from the corresponding glycoside. Depending on the monosaccharide that is removed, the b-glycosidase is named b-glucosidase (glucose), b-galactosidase (galactose), b-xylosidase (xylose), and so on. Frequently, the same b-glycosidase is able to hydrolyze several different monosaccharide residues from glycosides. In this case, b-glucosidase (EC 3.2.1.21) is used to name all enzymes that remove glucose efficiently. The active site of these enzymes is formed by subsites numbered from the glycosidic linkage to be broken with negative (towards the nonreducing end of the substrate) or positive (towards the reducing end of the substrate) integers (Davies et al., 1997). The nonreducing

monosaccharide residue binds at the glycone (1) subsite, whereas the rest of the molecule accommodates at the aglycone subsite, which actually may correspond to several monosaccharide residue-binding subsites. Some insects have three or four digestive b-glycosidases with different substrate specificity. In others only two of these enzymes are found, that are able to hydrolyze as many different b-glycosides as the other three or four enzymes together (Ferreira et al., 1998; Azevedo et al., 2003). Insect b-glycosidases best characterized have molecular masses of 30–150 kDa, pH optima of 4.5–6.5, and pI values of 3.7–6.8, whereas Km values with cellobiose or p-nitrophenyl b-glucoside (NpbGlu) as substrates, are found in the range of 0.2–2 mM. Although hydrolyzing several similar substances, insect digestive b-glycosidases have different specificities, preferring b-glucosides or b-galactosides as substrates, with hydrophobic or hydrophilic moieties in the aglycone part of the substrate (Terra and Ferreira, 1994; Azevedo et al., 2003). A few insect digestive b-glycosidases are more active on galactosides than on glucosides. Such enzymes are found in Locusta migratoria adults (Morgan, 1975), Abracris flavolineata adults (Marana, Terra, and Ferreira, unpublished data). In Rhynchosciara americana (Ferreira and Terra, 1983) and T. molitor (Ferreira et al., 2003) larvae, there is a b-glycosidase that hydrolyzes galactosides but not glucosides. Based on relative catalytical efficiency on several substrates, insect b-glycosidases can be divided into two classes. Class A includes the enzymes that efficiently hydrolyze substrates with hydrophilic aglycones, such as disaccharides and oligosaccharides. Class B comprises enzymes that have high activity only on substrates with hydrophobic aglycones, such as alkyl-, p-nitrophenyl-, and methylumbelliferyl-glycosides. Based on the same properties, b-glycosidases were previously divided into three classes (Terra and Ferreira, 1994), but the characterization of more enzymes showed that the previously proposed class 2 of b-glycosidases does not exist. Enzymes from the former class 2 comprised b-glycosidases supposed to have high activity only on di- and oligosaccharides. Former classes 2 plus 1 are now grouped in class A. Former class 3 are now named class B. Enzymes from class A are more abundant than b-glycosidases from class B. Class A b-glycosidases hydrolyze di- and oligosaccharides and have four subsites for glucose binding in the active site: one in the glycone (1) and three in the aglycone (þ1, þ2, þ3) position (Ferreira et al., 2001, 2003;

Biochemistry of Digestion

Marana et al., 2001; Azevedo et al., 2003). Some enzymes seem to be adapted to use disaccharides besides oligosaccharides as substrates. Optimal hydrolysis of disaccharides relies on high affinities to glucose moieties in 1 and primarily in the þ1 subsite (Ferreira et al., 2003). The enzymes highly active against oligosaccharides have subsites 1, þ1, and þ2 with similar affinities to glucose moieties (Ferreira et al., 2001, 2003). The affinities of some b-glycosidases for alkylglucosides were determined and binding energies for each methylene group to the active site were calculated. Values obtained for class A enzymes from Spodoptera frugiperda bGly50 (Marana et al., 2001) and T. molitor bGly1 (Ferreira et al., 2001) are 1.3 kJ mol1 and 0.97 kJ mol1, respectively. Surprisingly, the binding energy in the case of A. flavolineata alkyl-glycosidase, which is an enzyme from class B, is only 0.47 kJ mol1 (calculated from Marana et al., 1995). Since the enzymes from T. molitor and S. frugiperda can hydrolyze diand oligosaccharides and the A. flavolineata alkyl b-glycosidase can only use synthetic and more hydrophobic compounds as substrates, it seems that the hydrophobicity of the aglycone region is not directly related to the type of substrate that is hydrolyzed by the enzyme. Probably the extension of the aglycone site plays a major role. Class A b-glycosidases are able to hydrolyze b-1,3, b-1,4, and b-1,6 glycoside bonds from diand oligosaccharides. These enzymes are likely to be involved in the intermediate and terminal digestion of cellulose, hemicellulose, and glycoproteins present in food. Class B b-glycosidases such as D. saccharalis bGly2 (Azevedo et al., 2003) and T. molitor bGly4 seem to have two active sites (Ferreira et al., 2003). These were detected by using alternative substrates as inhibitors. Two active sites are also found in bGly47 from S. frugiperda (Marana et al., 2000) which is a class A enzyme, but one of its active site has properties of class B b-glycosidase. The three enzymes with two putative active sites are the main enzymes responsible for b-galactoside hydrolysis in the insect gut. The enzymes from the lepidopteran S. frugiperda and D. saccharalis hydrolyze mainly galactosides in one active site and glucosides in the other. The presence of two active sites in mammalian lactase-phlorizin hydrolase (the digestive mammalian b-glycosidase) has been known for a long time. The difference between the enzymes is that insect b-glycosidases with two active sites have the size of only one of the two mammalian b-glycosidase domains. Unfortunately, none of these types of enzymes has yet been cloned in

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insects. It is interesting to note that T. molitor bGly4 is activated by the detergent Triton X-100. Class B b-glycosidases (or active site) with high activity against hydrophobic substrates may have the physiological role of hydrolyzing glycolipids, mainly galactolipidis that are found in high amount in vegetal tissues. The main galactolipids in plants are 2,3-diacyl b-galactoside d-glycerol (mono galactosyl diglyceride) and 2,3-diacyl 1-(a-galactosyl 1,6 b-galactosyl)-d-glycerol (digalactosyl diglyceride) (Harwood, 1980). These enzymes may act directly against the monogalactosyl diglyceride or on digalactosyl diglyceride after the removal of one of the galactose residues by a-galactosidase. The activation by amphipatic substances (as Triton X-100; see above) may be a mechanism to maintain high enzyme activity only in the neighborhood of plant cell membranes undergoing digestion in the insect midgut. Those membranes are the source of glycolipid substrates and activating detergent-like molecules. Distant from membranes, the b-glycosidase would be less active, thus hydrolyzing plant glucosides (see below) ingested by the insect with decreased efficiency. In agreement with the hypothesis presented above, bGly47 from S. frugiperda can hydrolyze glycosylceramide, although with low activity (Marana et al., 2000). In mammals, sphingosine and ceramide hydrolysis are dependent on enzyme activation by proteins called saposins (Harzer et al., 2001). Given that genome sequences similar to saposins were found in Drosophila melanogaster, insect b-glycosidases active on glycolipids may also need the same kind of activators, and their absence in the assay reaction may explain why the activity against ceramides is low or not detected at all. Free energy relationships (Withers and Rupitz, 1990) were used to compare the specificity of insect b-glycosidase subsites (Azevedo et al., 2003). The enzymes with more similar active sites are Diatraea saccharalis bGly3 and T. molitor bGly1, followed by D. saccharalis bGly1 and T. molitor bGly3. Diatraea saccharalis bGly1 and bGly3 have active sites somewhat similar to the S. frugiperda bGly50K and T. molitor bGly1 with T. molitor bGly3. D. saccharalis bGly2, T. molitor bGly4 (Ferreira et al., 2001), and S. frugiperda bGly47K (Marana et al., 2000) are similar in some aspects. The latter three enzymes seem to have two active sites and are, in each insect, primarily responsible for b-galactoside digestion. Since each of the three D. saccharalis b-glycosidases has a counterpart in T. molitor, Azevedo et al. (2003) speculated that insects with the same number of b-glycosidases could have similar enzymes.

186 Biochemistry of Digestion

The only insect b-glycosidases sequenced to date have been those from the moth S. frugiperda midgut (bGly50) (Marana et al., 2001), the beetle T. molitor midgut (bGly1) (Ferreira et al., 2001), and termite Neotermes koshunensis salivary glands (Tokuda et al., 2002). The three enzymes share about 50% identity with one another and to other enzymes from glycosyl hydrolases from family 1, which have two glutamic acid residues responsible for catalysis and a structure of (b/a)8-barrel. In the bGly50 from S. frugiperda the pKa of the nucleophile (Glu 399) is 4.9 and of the proton donor (Glu 187) is 7.5. In this enzyme, residue Glu 451 seems to be a key residue in determining the enzyme preference for glucosides versus galactosides. This is due to its interaction, in the glycone site, with substrate equatorial or axial hydroxyl 4, which is the only position where glucose differs from galactose. The steric hindrance of the same residue with hydroxyl 6 probably also explains why fucosides are the best substrate for many b-glycosidases (Marana et al., 2002b). Besides having a role in digestion, b-glycosidases are important in insect–plant relationships. To avoid

herbivory plants synthesize a large number of toxic glucosides (Figure 6) which may be present in concentrations from 0.5% to 1% (Spencer, 1988). The presence of these glucosides in some insect diets may explain the variable number of b-glycosidases with different specificities present in their guts. Most plant glucosides have a hydrophobic aglycone and are b-linked O-glycosyl compounds. Since aglycones are usually more toxic than the glycosides themselves, intoxication may be avoided by decreasing the activity of the enzyme most active on toxic glucosides, without affecting the final digestion of hemicellulose and cellulose carried out by the other enzymes. This is exemplified by D. saccharalis larvae, which have three b-glycosidases in their midgut, feeding on diets containing the cyanogenic glucoside amygdalin. In this condition, the activity of the enzyme responsible for the hydrolysis of prunasin is decreased (Ferreira et al., 1997). Prunasin is the saccharide resulting after the removal from amygdalin, and that forms the cyanogenic mandelonitrille upon hydrolysis (Figure 6). Resistance to toxic glucosides may also be achieved by the lack of enzymes able to hydrolyze toxic b-glucosides,

Figure 6 b-Glucosidase acting on a cyanogenic glucoside releases glucose and cyanohydrin that spontaneously decomposes, producing a ketone (or an aldehyde) and hydrogen cyanide. If R1 ¼ R2 ¼ CH3, the glucoside is linamarin and the resulting ketone is acetone. If R1 ¼ H and R2 ¼ phenyl, the glucoside is prunasin and the resulting aldehyde is benzaldehyde (see more examples in Vetter (2000)).

Biochemistry of Digestion

as observed with S. frugiperda larvae, which have two b-glycosidases unable to efficiently hydrolyze prunasin (Marana et al., 2001; S.R. Marana, personal communication). Progress in this field will require disclosing the mechanisms by which the presence of toxic b-glucosides differentially affects the midgut b-glycosidases and knowing the details of the active site architecture responsible for the specificity of these enzymes. 4.5.4.8. Trehalases

Trehalase (EC 3.2.1.28) hydrolyzes a,a0 -trehalose into two glucose molecules and is one of the most widespread carbohydrases in insects, occurring in most tissues. Trehalase is very important for insect metabolism, since trehalose is the main circulating sugar in these organisms. Apical and basal trehalases can be distinguished in insect midguts. The apical trehalase may be soluble (glycocalyxassociated or secreted into the midgut lumen) or microvillar, whereas the midgut basal trehalase is an integral protein of the basal plasma membrane. The apical midgut trehalase is a true digestive enzyme. The midgut basal trehalase probably plays a role in the midgut utilization of hemolymph trehalose (Terra and Ferreira, 1994). In spite of the importance of trehalase, it is poorly studied in insects and also in other sources. There is not a single trehalase with known catalytical groups or with its 3D structure resolved. Trehalases partially or completely purified from insect guts have pH optima from 4.8 to 6.0, Km from 0.33 to 1.1 mM, pI around 4.6, and molecular masses from 60 to 138 kDa (Terra and Ferreira, 1994). Tris is usually a competitive inhibitor of trehalase. Inhibition of midgut trehalases at their optimum pH was reported in Apis mellifera, R. americana, and B. mori, with Kis of 50, 74, and 47 mM, respectively, (Terra and Ferreira, 1994). Tris inhibited R. americana trehalase at pH 9.0 with a higher Ki (182 mM) than at pH 6.0 (74 mM), suggesting the presence of a negative charge at the active site to which the protonated Tris binds (Terra et al., 1978). In the Lepidoptera Lymantria dispar (Valaitis and Bowers, 1993) and S. frugiperda (Silva, Terra, and Ferreira, unpublished data) Tris up to 100 mM does not inhibit the enzymes near the pH optimum. Contrasting to what was found in trehalase from R. americana, in S. frugiperda Tris is a competitive inhibitor with a small Ki (0.55 mM) at pH 9.0. The results may be related to differences in the active site of midgut trehalases from Lepidoptera. There have been a few attempts to identify important groups in the midgut trehalase active site. Terra

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et al. (1978, 1979, 1983) determined the pKa values of the catalytical groups of the R. americana midgut trehalase. The pKa value of the nucleophile was 5.0 (kinetic data) or 5.3 (carbodiimide modification results), whereas the pKa value of the proton donor was 8.3 (kinetic data) or 7.7 (carbodiimide modification). Since there was a disagreement between the pKa values determined for the proton donor, and taking into account that carbodiimide modification is only partially protected by trehalose, the authors suggested that the proton donor is near, but not at the active site, and that it participates in the reaction through another amino acid residue, like histidine (Terra et al., 1979). Lee et al. (2001) with the same approach as Terra et al. (1979) found pKa values of 5.3 and 8.5 for the A. mellifera trehalase. These authors and Valaitis and Bowers (1993), who worked with L. dispar trehalase, showed that the trehalases were inactivated by diethyl pyrocarbonate (DPC), but the work did not progress further. Purified midgut S. frugiperda trehalase is modified by DPC only in the presence of 2  Ki of a-methyl glucoside, a linear competitive inhibitor of the enzyme, indicating that there is a conformational change in the enzyme following inhibitor binding. The modification achieved with DPC affects an imidazole group and only inhibits 60% of the enzyme activity. The pKa values obtained from kinetic data are 4.8 and 7.6, and the pKa values calculated from carbodiimide and phenyl glyoxal modification are 4.9 and 7.8, respectively. Whereas 10 Km trehalose competitively protect the enzyme from phenyl glyoxal modification, only partial protection is seen when the modifier is carbodiimide. These and other results suggest that a carboxyl group is the nucleophile and an Arg residue is the proton donor, which pKa is affected by a nearby imidazole group (Silva, Terra, and Ferreira, unpublished data). The only insect midgut trehalase sequenced up till now is that from B. mori pupae (Su et al., 1993). The corresponding mRNA is actively transcribed in midgut and Malpighian tubules, but not in the other tissues (Su et al., 1994). It is interesting to note that, in mammals, the same trehalases are present in the microvillar membranes of small intestine and kidney. The midgut trehalase from B. mori has identity of only 46% with trehalase from the male accessory gland from T. molitor. Since similar low identities are found for B. mori trehalase (known to be from midgut) and the other insect trehalases deposited in the GenBank, these probably do not have a midgut origin. Plant toxic b-glucosides and their aglycones can inhibit, with varied efficiency, some or all trehalases

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from Malpighian tubules, fat body, midgut, and body wall of P. americana, M. domestica, S. frugiperda, and D. saccharalis (Silva, Terra, and Ferreira, unpublished data). Toxic b-glucosides are produced by many plant species and are present in high concentrations (see Section 4.5.4.7). It is not known whether those glucosides or aglycones are absorbed by the insect gut and interact with trehalases in tissues other than the midgut. It would be interesting to know if this happens and how insects resistant to toxic b-glucosides avoid the damage they can cause. 4.5.4.9. Acetylhexosaminidases, b-Fructosidases, and a-Galactosidases

An enzyme related to chitinolytic enzymes is b-Nacetyl-d-hexosaminidase (EC 3.2.1.52), which differs from b-N-acetyl-d-glucosaminidase in having a rather wide substrate specificity. The enzyme is found in many insects and its presumed physiological role is the hydrolysis of N-acetylglucosamine b-linked compounds such as glycoproteins (Terra and Ferreira, 1994). Detailed studies of this enzyme are lacking. Sucrose hydrolysis is catalyzed by enzymes that are specific for the a-glucosyl (a-glucosidase, EC 3.2.1.20; see above) or for the b-fructosyl residue (b-fructosidase, EC 3.2.1.26) of the substrate. b-Fructosidase is characterized by its activity toward sucrose and raffinose and lack of activity upon maltose and melibiose. In insect midguts, sucrose hydrolysis generally occurs by action of the conspicuous a-glucosidase rather than by b-fructosidase. Only a small number of reports verify the presence of b-fructosidase in insects (Terra and Ferreira, 1994; Scrivener et al., 1997) and there has been only one attempt (Santos and Terra, 1986a) at characterization. Larvae and adults of the moth Erinnyis ello have a midgut b-fructosidase with pH optimum 6.0, Km 30 mM (sucrose), pI 5.2, and molecular mass of 78 kDa. The physiological role of this enzyme is to hydrolyze sucrose, the major leaf (larvae) or nectar (adults) carbohydrate, which is not efficiently digested by E. ello midgut a-glucosidase (Santos and Terra, 1986b). a-Galactosidase (a-d-galactoside galactohydrolase, EC 3.2.1.22) catalyzes the hydrolysis of a-d-galactosidic linkages in the nonreducing end of oligosaccharides, galactomannans, and galactolipids and is widely distributed in nature (Dey and Pridham, 1972). Galactooligosaccharides, such as melibiose, raffinose, and stachiose are common in plants, mainly in lipid-rich seeds (Shiroya, 1963), whereas galactolipids are widespread among leaves. The major lipids in chloroplast membranes are monogalactosyldiglyceride and digalactosyldiglyceride (Harwood, 1980).

There have been few attempts to resolve insect midgut a-galactosidases. Gel filtration and heat inactivation suggested that there is a single a-galactosidase activity (30 kDa, pH optimum 5.0) in Dysdercus peruvianus midgut that is more efficient on raffinose than on melibiose and NPaGal (Silva and Terra, 1997). There are two a-galactosidases in Abracris flavolineata midguts: the major (112 kDa, pH optimum 5.4) is active on melibiose and raffinose in addition to NPaGal, whereas the minor (70 kDa, pH optimum 5.7) hydrolyzes only NPaGal (Ferreira et al., 1999). In the case of Psacothea hilaris, gel filtration gave evidence of the presence of multiple overlapping a-galactosidases more active on NPaGal than on melibiose (Scrivener et al., 1997). There are three midgut luminal a-galactosidases (TG1, TG2, and TG3) from T. molitor larvae that are partially resolved by ionexchange chromatography (Grossmann and Terra, 2001). The enzymes have approximately the same pH optimum (5.0), pI value (4.6), and molecular mass (46–49 kDa). TG2 hydrolyzes a-1,6-galactosaccharides, exemplified by raffinose, whereas TG3 acts on melibiose and apparently also on digalactosyldiglyceride, the most important compound in thylacoid membranes of chloroplast, converting it into monogalactosyldiglyceride. Spodoptera frugiperda larvae have three midgut a-galactosidases (SG1, SG2, and SG3) partially resolved by ionexchange chromatography (Grossmann and Terra, 2001). The enzymes have similar pH optimum (5.8), pI value (7.2), and molecular mass (46–52 kDa). SG1 and SG3 hydrolyze melibiose and SG3 digests raffinose and, perhaps, also digalactosyldiglyceride.

4.5.5. Digestion of Proteins 4.5.5.1. Initial Considerations

The initial digestion of proteins is carried out by proteinases (endopeptidases) that break internal bonds in proteins. Different proteinases are necessary to do this because the amino acid residues vary along the peptide chain. There are three subclasses of proteinases involved in digestion classified according to their active site group (and hence by their mechanism): serine, cysteine, and aspartic proteinases. In each of the mentioned subclasses, there are several proteinases differing in substrate specificities. The oligopeptides resulting from proteinase action are attacked from the N-terminal end by aminopeptidases and from the C-terminal end by carboxypeptidases, both enzymes liberating one amino acid residue at each catalytic step. Finally, the dipeptides formed are hydrolyzed by dipeptidases.

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4.5.5.2. Serine Proteinases

Serine proteinases (EC 3.4.21) (MEROPS) have serine, histidine, and aspartic acid residues (called the catalytic triad) in the active site. The family of enzymes homologous to chymotrypsin (Barrett et al., 1998) includes the major digestive enzymes trypsin, chymotrypsin, and elastase. These enzymes differ in structural features that are associated with their different substrate specificities, as detailed below. The numbering of residues in enzyme polypeptide chains is referred to that of bovine chymotrypsin. 4.5.5.2.1. Trypsins Trypsins (EC 3.4.21.4) preferentially cleave protein chains on the carboxyl side of basic l-amino acids such as arginine and lysine. Most insect trypsins have molecular masses in the range 20–35 kDa, pI values 4–5, and pH optima 8–10. These enzymes occur in the majority of the insects, with the remarkable exception of hemipteran species and some taxa belonging to the series Cucujiformia of Coleoptera like Curculionidae (Terra and Ferreira, 1994). Nevertheless, some heteropteran Hemiptera have trypsin in the salivary glands (Zeng et al., 2002). Trypsin is usually identified in insect midgut homogenates using benzoyl-arginine p-nitroanilide (B-R-pNA, often referred to as BApNA) or benzoylarginine 7-amino-4-methyl coumarin (B-R-MCA) as substrates and with N-a-tosyl-l-lysine chloromethyl ketone (TLCK), phenylmethylsulfonyl fluoride (PMSF), or diisopropylfluorophosphate (DFP) as inactivating compounds. The substrates of choice for assaying insect trypsins are those in Figure 7. Trypsins from Orthoptera, Dictyoptera, and Coleoptera are usually purified by ion-exchange chromatography, whereas those from Diptera and Lepidoptera, by affinity chromatography, either in benzamidineagarose (elution with benzamidine or by change in pH) or in soybean trypsin inhibitor (SBTI)-Sepharose (elution by change in pH). Due to significant autolysis, lepidopteran trypsins are more frequently purified by chromatography on benzamidine-Agarose with elution with benzamidine. A total of 109 insect trypsin sequences were registered in GenBank in March 2003, corresponding to 34 species pertaining to five orders. Examples may be found among Hemiptera (Zeng et al., 2002), Coleoptera (Girard and Jouanin, 1999a; Zhu and Baker, 1999), Diptera (Davis et al., 1985; Barillas-Mury et al., 1993), Siphonaptera (Gaines et al., 1999), and Lepidoptera (Peterson et al., 1994; Zhu et al., 2000). Several sequences were not complete and some insect species are represented by many sequences that probably include enzymes other than digestive enzymes. The complete sequences have signal and activation

Figure 7 Substrates used in the assay of enzymes involved in protein digestion. Sn are subsites in the enzymes and Pn are amino acid residues in substrates. The arrows point to bonds cleaved by the different enzymes. Abz-Xn-EDDnp is a class of peptides with quenching (EDDnp) and fluorescent (Abz) groups at the C- and N-terminal ends, respectively, so that after hydrolysis the peptides become fluorescent. Substrates with C-terminal MCA are fluorescent and those with pNA are colorimetric. Contrary to GL, LG is also hydrolyzed by APN in addition to dipeptidase. For further details, see text. TRY, trypsin; CHY, chymotrypsin; ELA, elastase; CAL, cathepsin L-like enzyme; ASP, aspartic proteinase; APN, aminopeptidase N; CPA, carboxypeptidase A; CPB carboxypeptidase B; DIP, dipeptidase.

peptides and the features typical of trypsin-like enzymes, including the conserved N-terminal residues IVGG, the catalytic amino acid triad of serine proteinase active sites (His 57, Asp 102, and Ser 195), three pairs of conserved cysteine residues for disulfide bonds, and the residue Asp 189 that determines specificity in trypsin-like enzymes (see Figure 5). In spite of having structural features resembling vertebrate trypsins, insect trypsins differ from these because they are not activated or stabilized by calcium ions and frequently are unstable in acidic pH (Terra and Ferreira, 1994). Another contrasting aspect is the almost lack of reports on the isolation of inactive insect trypsinogens, that is, trypsin molecules still having the activation peptide (Reeck et al., 1999). The activation of insect trypsinogen to trypsin deserves a closer look. Finally, other differences between vertebrate and insect trypsins include their substrate specificities and their interaction with protein inhibitors.

190 Biochemistry of Digestion

Amino acyl residues in proteinase substrates are numbered from the hydrolyzed peptide bonds as P1, P2, P3, . . . , Pn in the direction of the N-terminus and P0 1, P0 2, P0 3, . . . , P0 n in the direction of the peptide C-terminus, whereas the corresponding enzyme subsites are numbered S1, S2, S3, . . . , Sn and S0 1, S0 2, S0 3, . . . , S0 n (Schechter and Berger, 1967) (Figure 7). Mammalian trypsin preferably cleaves substrates having arginine rather than lysine at P1 (primary specificity) (Craik et al., 1985). The same primary specificity was found for insect trypsins, except those from lepidopterans, which prefer lysine at P1 (Lopes et al., 2004). This will be discussed below in relation to trypsin insensitivity to protein inhibitors. In order to characterize the trypsin specificity at subsites other than S1, quenched fluorescent substrates like o-aminobenzoyl-GGRGAGQ-2,4dinitrophenyl-ethylene diamine (where R stands for arginine at P1 position) were synthesized with 15 amino acid replacements at each of the positions P01 , P02 , P2, and P3. The results suggested that trypsin subsites are more hydrophobic in trypsins from the more evolved insects (A.R. Lopes et al., unpublished data). Trypsin from different insects also differ in the strength their subsites bind the substrate or the transition state (high-energy intermediate of the reaction). In other words, trypsin subsites differ in how they favor substrate binding or catalysis (Marana et al., 2002a). Plants have protein inhibitors (PIs) of insect midgut serine proteinases that affect insect development (Ryan, 1990). Insects may adapt to the presence of PIs in diet by overexpressing proteinases (Bown et al., 1997; Broadway, 1997; Gatehouse et al., 1997), by proteolytical inactivation of PIs mediated by the insect’s own proteinases (Giri et al., 1998), or by expressing new proteinases that are resistant to the inhibitor (Mazumbar-Leighton and Broadway, 2001a, 2001b). Current research is investigating the molecular basis of the difference between sensitive and inhibitor-insensitive trypsins, as well as the regulation of these enzymes. PIs produced by plants have a region, named the reactive site, that interacts with the active site of their target enzymes. The reactive sites of many PIs are hydrophilic loops with a lysine residue at P1 (Lopes et al., 2004). As lepidopteran trypsins have hydrophobic subsites and prefer lysine rather than arginine at P1 (see above), they are usually more resistant to PIs than the other insect trypsins. In this respect, it is interesting to note that PI-insensitive trypsins from Heliothis virescens bind more tightly to a hydrophobic chromatographic column than sensitive trypsins do (Brito et al., 2001). These observations lead to the hypothesis that the

molecular differences between sensitive and insensitive trypsins must rely on the interactions of PIs with residues in and around the enzyme active site. An interesting approach to study insect–PI interactions was introduced by Volpicella et al. (2003). They compared the sequence of a sensitive trypsin from Helicoverpa armigera with the insensitive trypsin from the closely related species H. zea. The 57 different amino acids observed between the two enzymes were superimposed on the porcine trypsin crystal structure, where the residues known to be in contact to a Kunitz-type inhibitor (Song and Suh, 1998) were identified. The residues at positions (chymotrypsin numbering) 41, 57, 60, 95, 99, 151, 175, 213, 217, and 220 were considered by Volpicella et al. (2003) to be important in H. zea trypsin–PI interaction. However, some of the interacting residues may have been misidentified because trypsins from different species were compared. In a similar approach, Lopes et al. (2004) aligned all available trypsin sequences characterized as sensitive or insensitive to Kunitz-type inhibitor (Bown et al., 1997; Mazumbar-Leighton and Broadway, 2001a) with porcine trypsin. After discounting conserved positions and positions not typical of sensitive or insensitive trypsins, the remaining positions that agree with those involved in porcine trypsin–PI (Bowman-Birk type, Lin et al., 1993; Kunitz type, Song and Suh, 1998) or substrate (Koepke et al., 2000) interactions were: 60, 94, 97, 98, 99, 188, 190, 213, 215, 217, 219, 228. These positions support the tree branches in a neighbor-joining analysis of sensitive (I, III) and insensitive (II) trypsin sequences (Lopes et al., 2004) (Figure 8a). Sitedirected mutagenesis of trypsin, followed by the determination of the binding constants of mutated trypsins with PIs, may help to resolve the discrepancy. The mechanism by which PIs in diet induces the synthesis of insensitive trypsin in responsive insects remains completely unknown, as well as the regulatory element that may be involved. 4.5.5.2.2. Chymotrypsins Chymotrypsin (EC 3.4.21.1) preferentially cleaves protein chains at the carboxyl side of aromatic amino acids. Insect chymotrypsins usually have molecular masses of 20–30 kDa and pH optima of 8–11, and they differ from their vertebrate counterparts in their instability at acidic pH, inhibition pattern with SBTI (Terra and Ferreira, 1994) and, finally, in reacting with N-a-tosyl-l-phenylalanine chloromethyl ketone (TPCK) (see below). The distribution of chymotrypsin among insect taxa is similar to that of trypsin (Terra and Ferreira, 1994), including the occurrence in the salivary glands of some heteropteran bugs

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191

Figure 8 Neighbor-joining distance analysis tree of insect trypsin and chymotrypsin sequences. (a) Clusters formed by sequences of Lepidoptera trypsins sensitive (I, III) or insensitive (II) to protein inhibitors according to Bown et al. (1997) and Mazumbar-Leighton and Broadway (2001b). The amino acid positions important in defining the clusters are indicated. (b) Clusters formed by sequences of insect chymotrypsins. The sequences are identified by two digits: the first denotes the insect species (see below) and the second identify the different sequences of the same insect. Trypsin sequences: 1, Agrotis ipsilon; 2, Helicoverpa zea; 3, H. armigera. Chymotrypsin sequences: 1, Plodia interpunctella; 2, H. armigera; 3, H. zea; 4, A. ipsilon; 5, Spodoptera frugiperda; 6, Manduca sexta; 7, Heliothis virescens; 8, Phaedon cochleariae; 9, Scirpophaga incertulas; 10, Anopheles gambiae; 11, A. aquasalis; 12, A. darlingi; 13, Rhyzopertha dominica; 14, Ctenocephalides felis; 15, Vespa crabro; 16, V. orientalis; 17, Solenopsis invicta; 18, Aedes aegypti; 19, Culex pipiens pallens; 20, Glossina morsitans morsitans; 21, cow; 22, rat. (Courtesy of A.R. Lopes.)

(Colebatch et al., 2002). The earlier failure to detect chymotrypsin activity in insect midguts was a consequence of using the mammalian chymotrypsin substrates, like benzoyl-tyrosine p-nitroanilide (B-Y-pNA) or benzoyl-tyrosine ethyl ester (B-Y-ee), in the assays. Insect chymotrypsins have an extended active site and larger substrates, like succinyl-AAPF-p-nitroanilide (Suc-AAPF-pNA), are necessary for their detection (Lee and Anstee, 1995; Lopes et al., 2004) (Figure 7). Insect chymotrypsins are usually purified by affinity chromatography in phenyl butylamina-Sepharose (elution with

phenyl butylamina) or in SBTI-Sepharose (elution with benzamidine) for enymes from lepidopterans, and by ion-exchange chromatography for those from dictyopterans, orthopterans, hymenopterans, and dipterans. There are 63 complete sequences of insect chymotrypsins registered in the GenBank (as of March 2003) corresponding to 22 species pertaining to six different orders. Examples are found among Hemiptera (Colebatch et al., 2002), Coleoptera (Girard and Jouanin, 1999a; Zhu and Baker, 2000), Hymenoptera (Jany et al., 1983; Whitworth et al., 1998),

192 Biochemistry of Digestion

Siphonaptera (Gaines et al., 1999), Diptera (Casu et al., 1994; de Almeida et al., 2003), and Lepidoptera (Peterson et al., 1995; Bown et al., 1997). All the sequences have a signal peptide, an activation peptide (ending with an arginine residue), the catalytic triad (His 57, Asp 102, and Ser 195), three pairs of conserved cysteine residue, conserved N-terminal sequence IVGG and Ser/Gly/Tyr 189 which confers specificity to chymotrypsin-like enzymes (Figure 5). The insect digestive chymotrypsin that has been most thoroughly studied is that of Manduca sexta (Lepidoptera: Sphingidae) (Peterson et al., 1995). In this enzyme, the activation peptide is longer and has a net charge different from that of bovine chymotrypsinogen, leading the authors to suggest that the insect enzyme is activated by a peculiar mechanism. The mammalian chymotrypsin has a pH optimum around 8 with two catalytic important pKas of 6.8 and 9.5, corresponding to the active-site histidine, and N-terminal leucine, respectively. In contrast, the M. sexta chymotrypsin has pH optimum 10.5–11 and a single kinetically significant pKa at pH 9.2. This pKa may represent the active-site histidine in an appropriate environment, although several other hypotheses were discussed (Peterson et al., 1995). It is not clear whether the insect chymotrypsin active-site changes associated with TPCK resistance (see below) may also be the cause of the putative histidine pKa displacement. The resolution of the 3D structure of the fire ant digestive chymotrypsin led to the conclusion that it is strikingly similar to mammalian chymotrypsin, but has differences beyond those found among homologs from different mammalian systems (Botos et al., 2000). The similarities include a conserved backbone scaffold and structural domains. Differences include the activation mechanism and substitutions in the subsite S1 and mainly in the other subsites (S4–S04 ) that suggest different substrate specificities and interactions with PIs. In agreement with this, different insect chymotrypsins

are sensitive to distinct PIs and like trypsins, PI-insensitive chymotrypsins may be induced in insects ingesting PI-containing diets (Bown et al., 1997; Mazumbar-Leighton and Broadway, 2001b). Chymotrypsin sequences form several branches in a neighbor-joining analysis that correspond to phylogenetic groups (Figure 8b). Lepidoptera species are clustered in the two branches supported by Trp 59 (all sequences have a conserved tryptophan at position 59), except for the oligophagous pyralid Scirpophaga incertulas (9 in Figure 8b). One of the major lepidopteran branches corresponds to sequences from the pyralid Plodia interpunctella (1 in Figure 8b) and the other two sequences from noctuids and sphingids, all of them polyphagous. The data suggest that Trp 59 is related to a polyphagous habit. Chymotrypsins from insects that routinely ingest ketone-releasing compounds (like several plant glycosides) (see Figures 6 and 9) are not affected much by these compounds and others that react with His 57. Thus, in comparison with bovine chymotrypsin, the chymotrypsin from polyphagous lepidopteran insects reacts slowly with chloromethyl ketones, whereas those of oligophagous pyralid insects react rapidly (A.R. Lopes et al., unpublished data). Modeling Spodoptera frugiperda (Noctuidae) chymotrypsin, based on its sequence and on crystallographic data of bovine chymotrypsin, showed that His 57 is probably protected from alkylation by a bulky (tryptophan) residue at position 59 (A.R. Lopes et al., unpublished data). Bovine chymotrypsin has at position 59 Gly, thus rendering His 57 exposed to the medium. These adaptations are new examples of the interplay between insects and plants during their evolutionary arms race and deserve more attention through site-directed mutagenesis of recombinant chymotrypsins. 4.5.5.2.3. Elastases Since Christeller et al. (1990) described an elastase (EC 3.4.21.36)-like enzyme in

Figure 9 Ketones or aldehydes formed after the action of b-glucosidases on cyanogenic glucosides (Figure 6) may react with His 57 of chymotrypsin, inactivating it.

Biochemistry of Digestion

the cricket Teleogryllus commodus, this enzyme has been described in many other insects, including in homogeneous form (Terra and Ferreira, 1994; Whitworth et al., 1998). Usually elastase is identified with the substrate Suc-AAPL-pNA (Figure 7), combined with the observation of lack of activity on B-YpNA or B-Y-ee and resistance to TPCK. Since the mentioned substrate may also be hydrolyzed by chymotrypsin and lack of activity on B-Y-pNA and/or resistance to TPCK are usual among chymotrypsins (see Section 4.5.5.2.2), most described elastase may actually be chymotrypsins. True elastases were isolated from gypsy moth midguts (Valaitis, 1995) and from whole larvae of Solenopsis invicta (Whitworth et al., 1998). The last-mentioned enzymes hydrolyze Suc-APA-pNA, but not substrates with phenylalanine at P1. Although the specific substrate for elastase (Suc-AAA-pNA) (Bieth et al., 1974) was not tested, the hydrolysis of Suc-AAAPV-pNA and the lack of hydrolysis of substrates with phenylalanine in P1 discount a chymotrypsin. One of the S. invicta elastases (E2) was cloned, sequenced, and shown to be more similar to chymotrypsin than to elastase (Whitworth et al., 1999). This work confirms the occurrence of elastase in insect midgut. Further work is necessary to evaluate the extent of this enzyme in insect midguts and the importance in digestion. 4.5.5.3. Cysteine Proteinases

Cysteine proteinase is usually assayed in insect midgut contents or midgut homogenates at pH 5–6 with B-R-pNA, B-R-NA, casein, or hemoglobin as substrate. Activation by sulfhydryl agents (dithiothreitol (DTT) or cysteine) and inhibition by transepoxysuccinyl-l-leucyl-amido (4-guanidinobutane) (E-64) are usually indicative of the presence of the enzyme. The observation of inhibition of hydrolytic activity on any of the mentioned substrates by E-64 is insufficient for a positive identification of cysteine proteinase. Trypsin hydrolyzes the same substrates and may be reversibly inhibited by E-64 (Novillo et al., 1997). The identification of cysteine proteinase was made easier with the substrate e-aminocaproyl-leucyl-(S-benzyl)-cysteinyl-MCA, which is hydrolyzed by cysteine proteinase but not by serine proteinases (Alves et al., 1996). Using such criteria, cysteine proteinases were described in Hemiptera Heteroptera and in species belonging to the series Cucujiformia of Coleoptera (Terra and Ferreira, 1994). Exceptions to this rule are the identification of cysteine proteinase in Hemiptera Auchenorrhyncha (aphids) (Cristofoletti et al., 2003) and the lack of this enzyme in cucujiform cerambycid beetles (Johnson and Rabosky, 2000).

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Insect midgut cysteine proteinases were at first denoted as cathepsin B (EC 3.4.22.1)-like enzymes, because cathepsin B was the first animal cysteine proteinase described. Later on it became known that cathepsin B is more important as a peptidyl dipeptidase, rather than as an endopeptidase, because of the existence of an extended loop that forms a cap to the active-site cleft, and carries a pair of histidine residues that are thought to bind to the C-terminal carboxylate of the substrate (Barrett et al., 1998). Cathepsin L (EC 3.4.22.15) is a true endopeptidase that preferentially cleaves peptide bonds with hydrophobic amino acid residues in P2 (cathepsin B prefers arginine at the same position) (Barrett et al., 1998). Thus, by using substrates like carbobenzoxy (Z)-FR-MCA and Z-RR-MCA it is possible to distinguish between the two enzymes (see Figure 7). Current research revealed that cathepsin L-like enzymes are the only insect cysteine proteinase quantitatively important. Much more difficult to ascertain is that a cathepsin L-like enzyme assayed in insect midguts has been secreted into midgut contents, and hence may be considered as a truly digestive enzyme. As in other animals (Barrett et al., 1998), cathepsin L-like enzymes in insects are expected to occur in lysosomes and never leave the cells. The same difficulties arise in trying to relate digestion to cathepsin L-like enzymes encoded by cDNAs cloned from midgut cells. The problems that may arise during cathepsin L-like enzyme characterization are well illustrated in a study with T. molitor larvae. Three cathepsin L-like sequences were recognized in a cDNA library prepared from T. molitor midguts. One sequence after being expressed and used to raise antibodies was found to correspond to a lysosomal cathepsin L immunolocalized mainly at hemocytes and fat body cells. The second sequence was not related yet with any enzyme active in midgut. Finally, the third one corresponds to a cathepsin L-like enzyme purified from midgut contents (Cristofoletti et al., unpublished data). Digestive cathepsin L-like enzymes have been purified to homogeneity only from Diabrotica virgifera (Coleoptera: Cucujiformia) (Koiwa et al., 2000), Acyrthosiphon pisum (Hemiptera: Auchenorrhyncha) (Cristofoletti et al., 2003), and T. molitor (Coleoptera: Cucujiformia) (Cristofoletti et al., unpublished data). The A. pisum enzyme is cell membrane-bound and faces the luminal contents, whereas D. virgifera and T. molitor ones are soluble enzymes secreted into midgut contents. The complete purification of those enzymes was achieved by affinity chromatography with soyacystatin as a ligand or by a combination of ion-exchange

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Figure 10 Neighbor-joining distance analysis tree of insect cathepsin L and aminopeptidase N sequences. (a) Clusters formed by cathepsin L-like sequences of insects known to have digestive cathepsins. (b) Clusters formed by coleopteran and lepidopteran aminopeptidases N. Sequences are identified as described in Figure 8. Cathepsin L-like sequences: 1, Myzus persicae; 2, Aphis gossypii; 3, Sitophilus zeamais; 4, Tenebrio molitor; 5, Rhodnius prolixus; 6, Diabrotica virgifera; 7, Phaedon cochleariae; 8, Hypera postica. Aminopeptidase N sequences: 1, Epiphyas postvittana; 2, Lymantria dispar ; 3, Bombyx mori; 4, Manduca sexta; 5, Helicoverpa armigera; 6, Heliothis punctigera; 7, H. virescens; 8, Plutella xylostella; 9, Plodia interpunctella; 10, Spodoptera littura; 11, Tenebrio molitor. (Courtesy of P. T. Cristofoletti.)

chromatographies. The enzymes have pH optima of 5–6, molecular masses of 20–40 kDa, prefer Z-FR-MCA over Z-RR-MCA, are inhibited by E-64, and activated by cysteine or DTT. At least the A. pisum enzyme is also inhibited by chymostatin (completely) and elastatinal (partly) (Cristofoletti et al., 2003). Twelve cathepsin L-like sequences corresponding to eight species of coleopterans and hemipterans (those known to have digestive cathepsins) are registered in GenBank (as of March 2003). The sequences demonstrate the features characteristic of family 1 of cysteine proteinases (Barrett et al., 1998): N-terminal propeptide that must be removed to activate the enzyme and the catalytic triad, Cys 25, His 169, and Asn 175 (papain numbering) and the ERFININ motif (Figure 5). The sequences form two

major branches in a neighbor-joining analysis (Figure 10a) (P.T. Cristofoletti et al., unpublished data). The data suggest that the Hemiptera digestive cathepsin is close to the lysosomal one from T. molitor (sequence 1), whereas those from the coleopterans (except from Sitophilis oryzae) are similar to the T. molitor digestive enzyme (sequence 3) (Figure 10a). It is probable that the S. oryzae enzymes are not true digestive but lysosomal, like T. molitor sequence 1 (P.T. Cristofoletti et al., unpublished data). More work is needed to clarify the role of cathepsin L-like enzymes in insect digestion. 4.5.5.4. Aspartic Proteinases

Aspartic proteinases are active at acid pH, hydrolyze internal peptide bonds in proteins, and attack some synthetic substrates, either chromophoric (Dunn

Biochemistry of Digestion

et al., 1986) or internally quenched fluorescent substrates (Pimenta et al., 2001) (Figure 7). Mammalian cathepsin D has a substrate-binding cleft that can accommodate up to seven amino acids and prefers to cleave between two hydrophobic residues (Barrett et al., 1998). The first report of aspartic proteinases in insects was made by Greenberg and Paretsky (1955), who found a strong proteolytic activity at pH 2.5–3.0 in homogenates of whole bodies of Musca domestica. Lemos and Terra (1991b) showed that the enzyme occurs in midguts and is cathepsin D-like. An aspartic proteinase similar to cathepsin D was found in families of Hemiptera, Heteroptera and in several families belonging to the cucujiform series of Coleoptera (Terra and Ferreira, 1994). Thus, it is possible that aspartic proteinases occur together with cysteine proteinase in Hemiptera and in most Coleoptera. The aspartic proteinase isolated from Callosobruchus maculatus (pH optimum 3.3, 62 kDa) (Silva and Xavier-Filho, 1991) and Tribolium castaneum (pH optimum 3.0, 22 kDa) (Blanco-Labra et al., 1996) were partially purified and shown to be similar to cathepsin D. These studies need to be extended, so that the origin, specificity, and structure of insect cathepsin D-like enzymes be clarified. 4.5.5.5. Aminopeptidases

Aminopeptidases sequentially remove amino acids from the N-terminus of peptides and are classified on the basis of their dependence on metal ions (usually Zn2þ or Mn2þ) and substrate specificity. Aminopeptidase N (EC 3.4.11.2) has a broad specificity, although it removes preferentially alanine and leucine residues from peptides, whereas aminopeptidase A (EC 3.4.11.7) prefers aspartyl (or glutamyl)peptides as substrates. Both are metalloenzymes (Nore´ n et al., 1986). In insect midguts, major amounts of soluble aminopeptidases are found in less evolved insects (e.g., Orthoptera, Hemiptera, Coleoptera Adephaga), whereas in more evolved insects (e.g., Coleoptera Polyphaga, Diptera, and Lepidoptera) aminopeptidase is found mainly bound to the microvillar membranes of midgut cells (Terra and Ferreira, 1994). Insect midgut aminopeptidases are metalloenzymes (ethylenediaminetetraacetic acid (EDTA) inhibition) and have pH optima of 7.2–9.0, irrespective of the pH of the midgut lumen from the different species, Km values (L-pNA) of 0.13–0.78 mM and molecular masses of 90–130 kDa. With a single exception (see below), all known insect aminopeptidases have a broad specificity, hydrolyzing a variety of amino acyl b-naphthylamides (except acidic amino acyl

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b-naphthylamides), indicating they are aminopeptidases N (Terra and Ferreira, 1994) (Figure 7). The exception is a soluble glycocalyx-associated midgut aminopeptidase from R. americana. This enzyme is an aminopeptidase removing N-terminal aspartic acid or glutamic acid residues from peptides that are not efficiently attacked by the other aminopeptidases (Klinkowstrom et al., 1994). In addition to a midgut aminopeptidase A, the dipteran R. americana has three midgut aminopeptidases N (one soluble and two membrane-bound). The soluble aminopeptidase N (115.7 kDa) prefers tetrapeptides over tripeptides (Ferreira and Terra, 1984), like the minor 107 kDa membrane-bound enzyme, whereas the contrary is true for the major 169 kDa membrane-bound aminopeptidase (Ferreira and Terra, 1985, 1986a, 1986b). The single midgut aminopeptidase N of the coleopterans Attagenus megatoma (Baker and Woo, 1981) and Tenebrio molitor (Cristofoletti and Terra, 1999) resemble the 115.7 kDa and 107 kDa aminopeptidase of R. americana. Approximately the same substrate specificity was observed with the two midgut aminopeptidases of the lepidopteran Tineola bisselliella (Ward, 1975a, 1975b). The data suggest that panorpoid insects (Diptera and Lepidoptera) present multiple aminopeptidases with different substrate specificities, in contrast with the single aminopeptidase of coleopterans. However, much more data are needed to support this hypothesis. There have been few attempts to characterize the active site of insect midgut aminopeptidases. Using multiple inhibition analysis and observing the protection against EDTA inactivation that different competitive inhibitors conferred to the enzyme, two subsites were proposed to occur in the active center of R. americana microvillar aminopeptidase: a hydrophobic subsite, to which isoamyl alcohol binds exposing the metal ion, and a polar subsite, to which hydroxylamine binds. Exposure of the metal ion after isoamyl alcohol binding may be analogous to the situation that results when part of the substrate occupies the hydrophobic subsite, causing conformational changes associated with the catalytic step (Ferreira and Terra, 1986b). The effect of pH at different temperatures on kinetic parameters of T. molitor midgut aminopeptidase and its inactivation by different compounds were studied (Cristofoletti and Terra, 2000). The data showed that T. molitor aminopeptidase catalysis depends on a metal ion, a carboxylate, and a protonated imidazole group and is, somehow, influenced by an arginine residue in the neighborhood of the active site. The catalytic metal binding depends on at least a deprotonated imidazole. In addition to the

196 Biochemistry of Digestion

above-mentioned groups involved in catalysis, at least one phenol group and one carboxylate are associated with substrate binding. Thus, T. molitor aminopeptidase shares common features with those of other zinc metallopeptidases, especially with mammalian aminopeptidase N, but it differs in some details. An imidazole group seems to be involved in catalysis in T. molitor aminopeptidase; this is not observed in mammalian aminopeptidase N, which has an imidazole group participating in substrate binding. Aminopeptidase N sequences are available for the following lepidopterans: Epiphyas postvittana, B. mori, Heliothis virescens, H. punctigera, Helicoverpa armigera, L. dispar, Manduca sexta, P. interpuctella, and Plutella xylostella (Gill et al., 1995; Knight et al., 1995; Denolf et al., 1997; Hua et al., 1998; Chang et al., 1999; Oltean et al., 1999; Yaoi et al., 1999; Garner et al., 1999; Emmerling et al., 2001; Rajagopal et al., 2003). The sequences have a signal peptide, a conserved RLP motif near the N-terminal, a zinc binding/gluzincin motif HEXXHX18E, a GAMEN conserved motif and a long hydrophobic C-terminal containing a glycosyl phosphatidyl inositol anchor (Figure 5). Based on the crystal structure of leukotriene A4 hydrolase, the two histidine residues and the distant glutamic acid residue of the gluzincin motif are zinc ligands, the glutamic acid residue between the histidine residues is involved in catalysis (Hooper, 1994; Rawlings and Barrett, 1995), and the glutamic acid residue of the GAMEN motif binds the substrate N-terminal amino acid (Luciani et al., 1998). In contrast to the situation in mammals, insect aminopeptidase N is membrane bound at the C-terminal. No soluble insect aminopeptidase N has been sequenced. Dendrograms derived from alignments of coleopteran and lepidopteran midgut aminopeptidases suggest that there are at least four groups of lepidopteran aminopeptidases, with the isoforms of the same animal distributed among the groups (Chang et al., 1999; Emmerling et al., 2001; Nakanishi et al., 2002; Rajagopal et al., 2003; P. T. Cristofoletti et al., unpublished data) (Figure 10b, clusters A, B, C, and D). The existence of a number of different aminopeptidases in lepidopterans could be explained by the need for enzymes with different substrate specificities (as shown above for R. americana) or different susceptibilities to inhibitors, similar to serine proteinases (see Section 4.5.5.2). Probably associated with the fact that aminopeptidases are major proteins in some microvillar membranes (55% of T. molitor midgut microvillar proteins) (Cristofoletti and Terra, 1999), they are targets of insecticidal Bacillus thuringiensis

crystal d-endotoxins. These toxins, after binding to aminopeptidases and receptor molecules called cadherins, form channels through which cell contents leak leading to death of the insect (Gill et al., 1995; Knight et al., 1995; Denolf et al., 1997). Although data on substrate specificity for lepidopteran aminopeptidase isoforms are lacking, there is evidence that the isoforms may have differences in toxin binding (Valaitis et al., 1997; Zhu et al., 2000; Nakanishi et al., 2002; Rajagopal et al., 2003). Cloning and sequencing dipteran aminopeptidases, for which differences in substrate specificity are known, and a study of substrate specificities of lepidopteran aminopeptidases, may clarify the selective advantages of the evolution of aminopeptidase groups. Furthermore, this study may support the hypothesis that aminopeptidase gene duplications have occurred in the panorpoid ancestor, before differentiation between dipterans and lepidopterans. 4.5.5.6. Carboxypeptidases and Dipeptidases

Carboxypeptidases hydrolyze single amino acids from the C-terminus of the peptide chain and are divided into classes on the basis of their catalytic mechanism. There are two digestive metallocarboxypeptidases in mammals: carboxypeptidase A (EC 3.4.17.1), which hydrolyzes, in alkaline medium, C-terminal amino acids, except arginine, lysine, and proline, and carboxypeptidase B (EC 3.4.1.7.2), which releases, in alkaline conditions, C-terminal lysine and arginine preferentially. Insect digestive carboxypeptidases have been classified as carboxypeptidase A or B depending on activity in alkaline medium against Z-GF (or hippuryl bphenyllactic acid) or Z-GR (or hippuryl-l-arginine), respectively (Figure 7). Digestive insect carboxypeptidase A-like enzymes are widespread among insects and most of them have pH otima of 7.5–9.0 and molecular masses of 20–50 kDa (Terra and Ferreira, 1994). They have been cloned and sequenced from Diptera (Ramos et al., 1993; Edwards et al., 1997) and Lepidoptera (Bown et al., 1998) and the enzyme from the lepidopteran H. armigera was also submitted to crystallographic studies (Este´ banez-Perpin˜ a´ et al., 2001). The sequences have signal and activation peptides and the features typical of carboxypeptidases A, including the residues His 69, Glu 72, and His 196, which bind the catalytic zinc ion, and Arg 71, Asn 144, Arg 145, and Tyr 248 responsible for substrate binding and Arg 127 and Glu 270 for catalysis. In spite of the overall similarity of H. armigera procarboxypeptidase with human procarboxypeptidase A2, there are differences in the loops between the conserved secondary structures, including the loop where the activation processing occurs.

Biochemistry of Digestion

Another important difference is the residue 255 (bottom of the S0 1 pocket) that defines the enzyme specificity. In mammalian sequences Asp 255 is found in carboxypeptidase B and Ile 255 in carboxypeptidase A. In insect carboxypeptidases A, this residue varies (but never is an acid residue) (Figure 5). Carboxypeptidases B-like enzymes have been detected in insect midguts (Terra and Ferreira, 1994) but none has been characterized in detail, because they are much less active than carboxypeptidases A. Dipeptidases hydrolyze dipeptides and are classified according to their substrate specificities. Dipeptidases comprise the poorest known of the insect peptide hydrolases. There have been few studies in which dipeptidase assays were performed and even fewer attempts to characterize the enzymes (Terra and Ferreira, 1994). The larval midgut of R. americana has three dipeptidases (two soluble with 63 kDa and 73 kDa, respectively, and one membrane-bound) that hydrolyze Gly-Leu, resembling dipeptide hydrolase (dipeptidase, E.C. 3.4.13.18), although in contrast to the mammalian enzyme they are very active upon Pro-Gly (Figure 7). Rhynchosciara americana also seems to have an amino acyl-histidine dipeptidase (carnosinase, EC 3.4.13.3) (Klinkowstrom et al., 1995). More work on insect digestive dipeptidases is urgently needed.

4.5.6. Digestion of Lipids and Phosphates 4.5.6.1. Overview

Lipids that contain fatty acids comprise storage lipids and membrane lipids. Storage lipids, such as oils present in seeds and fats in adipose tissue of animals, are triacylglycerols (triglycerides) and are hydrolyzed by lipases. Membrane lipids include phospholipids and glycolipids like mono- and digalactosyldiglycerides (see Section 4.5.4.7). Phospholipids are digested by phospholipases. A combination of a- and b-galactosidases may remove galactose residues from mono- and digalactosyldiglyceride to leave a diacylglycerol which may be hydrolyzed by a triacylglycerol lipase. Phosphate moieties need to be removed from phosphorylated compounds prior to absorption. This is accomplished by nonspecific phosphatases. The phosphatases may be active in an alkaline (alkaline phosphatase, EC 3.1.3.1) or acid (acid phosphates, EC 3.1.3.2) medium. 4.5.6.2. Lipases

Triacylglycerol lipases (EC 3.1.1.3) are enzymes that preferentially hydrolyze the outer links of

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triacylglycerols and act only on the water–lipid interface. Activity of the lipase is increased as the interface becomes larger due to lipid emulsification caused by emulsifiers (surfactants). Insects lack emulsifiers comparable to the bile salts of vertebrates, but surfactant phospholipids, including lysolecithin, occur in their midguts in sufficient concentration to alter the surface tension of midgut contents (De Veau and Schultz, 1992). Lysolecithin, and other surfactants, may be formed by the action of phospholipase A on ingested phospholipids (see below and Figure 1). Current research on insect lipases is focused on triacylglycerol lipase from the fat body and its interplay with flight muscles (Ryan and Van der Horst, 2000). Insect midgut triacylglycerol lipases have been studied in few insects and only in crude preparations. The data suggested that the enzyme preferentially releases fatty acids from the a-positions, prefers unsaturated fatty acids and is activated by calcium ions, thus resembling the action of mammalian pancreatic lipase. The resulting 2-monoacylglycerol may be absorbed or hydrolyzed (Terra et al., 1996). Hydrolysis of 2-monoacylglycerol may be accomplished by the triacylglycerol lipase, following migration of the fatty acid to the 1-position, which seems to be favored by the alkaline midgut pH, at least in M. sexta (Tsuchida and Wells, 1988). Esterases, which are usually named the carboxylesterases (ali-esterases, EC 3.1.1.1) catalyze the hydrolysis of carboxyl ester into alcohol and carboxylate. This enzyme, in contrast to lipases, attacks molecules that are completely dissolved in water. It also hydrolyzes water-insoluble long-chain fatty acid esters in the presence of surfactants, but at a rate much slower than that of triacylglycerol lipase. A role for esterases in digestion is unclear and because of this they are not reviewed in detail here. In spite of the fact that a requirement for essential fatty acids is probably universal in insects, progress has been limited in the study of lipid digestion. Presumably, the lack of comparatively simple, sensitive assays and the complexities of digestion related to lipid solubilization have hindered work in this area. Another reason to study enzymes associated with lipid digestion is that they might be important in limiting the development of pathogens and parasites. Hydrolysis of membrane lipids might cause cellular lysis and fatty acid products of digestion may possess antibiotic effects. 4.5.6.3. Phospholipases

Phospholipase A2 (EC 3.1.1.4) and phospholipase A1 (EC 3.1.1.32) remove from phosphatides the fatty acid attached to the 2-position and 1-position,

198 Biochemistry of Digestion

respectively, resulting in a lysophosphatide (Figure 2). Lysophosphatide is more stable in micellar aggregates than on membranes. Thus, the action of phospholipase A on the membrane phosphatides causes the solubilization of cell membranes, rendering the cell contents free to be acted upon by the appropriate digestive enzymes. Phospholipase is widespread among insects (Terra et al., 1996; Nor Aliza et al., 1999). Phospholipase A2 partially purified from the midgut of adult beetle Cincindella circumpicta has a molecular mass of 22 kDa and pH optimum 9.0, is calcium dependent, and is inhibited by the sitespecific inhibitor oleyoxyethyl phosphorylcholine. Unfed beetles did not express the phospholipase in the midgut contents (Uscian et al., 1995). Although lysophosphatide may be further hydrolyzed by a lysophospholipase (phospholipase B, EC 3.1.1.5), evidence suggests it is absorbed intact by insects (Turunen and Kastari, 1979; Weiher and Komnick, 1997). Phosphatides may also be hydrolyzed by phospholipase C (EC 3.1.4.3) yielding the phosphoryl base moiety and diacylglycerol, or by phospholipase D (EC 3.1.4.4), resulting phosphatidate and the base (Figure 2). Both enzymes have been found in insect midgut (Turunen, 1993), but have not studied in detail. 4.5.6.4. Phosphatases

Alkaline phosphatase is usually a midgut microvillar membrane marker in dipteran and lepidopteran species, although it may also occur in midgut basolateral membranes and even as a secretory enzyme. Acid phosphatase is usually soluble in the cytosol of midgut cells in many insects and may also appear in midgut contents or be found membrane-bound in midgut cells (Terra and Ferreira, 1994). The best-known alkaline phosphatases are those from B. mori (Lepidoptera: Bombycidae) larval midgut. The major membrane-bound and the minor soluble alkaline phosphatases were purified and shown to be monomeric enzymes with the following properties: (1) soluble enzyme, molecular mass of 61 kDa, pH optimum 9.8; (2) membranebound enzyme, molecular mass of 58 kDa, pH optimum 10.9. Both enzymes have wide substrate specificity and are inhibited by cysteine. The membrane-bound alkaline phosphatase occurs in the microvillar membranes of columnar cells, whereas the soluble enzyme is loosely attached to the goblet cell apical membrane facing the cell cavity (Eguchi, 1995). The determination of the complete sequence of the membrane-bound alkaline phosphatase led to the finding of putative regions for phosphatidylinositol anchoring, zinc-binding site but not for

N-glycosylation, despite the fact that the enzyme contains N-linked oligosaccharides (Itoh et al., 1991). The sequence of the soluble alkaline phosphatase was also determined and has high identity with the membrane-bound enzyme (Itoh et al., 1999). Acid phosphatases have been characterized in some detail only in Rhodnius prolixus (Hemiptera: Reduvidae). The major enzyme activity is soluble and has the following properties: wide specificity, a molecular mass of 82 kDa, Km for p-nitrophenyl phosphate 0.7 mM, and is inhibited by fluoride, tartrate, and molybdate. The minor enzyme activity is membrane-bound and is resolved into two enzymes (123 and 164 kDa) which are resistant to fluoride and tartrate (Terra et al., 1988).

4.5.7. The Peritrophic Membrane 4.5.7.1. The Origin, Structure, and Formation of the Peritrophic Membrane

There is a film surrounding the food bolus in most insects that occasionally is fluid (peritrophic gel) but more frequently is membranous (peritrophic membrane, PM). The PM is made up of a matrix of proteins (peritrophins) and chitin to which other components (e.g., enzymes, food molecules) may associate (see Chapter 4.3). This anatomical structure is sometimes called peritrophic matrix, but this term should be avoided because it does not convey the idea of a film and suggests it is the fundamental substance of some structure, usually filling a space as the mitochondrial matrix. The argument that membrane means a lipid bilayer is not valid because the PM is not a cell part, but an anatomical structure, like the nictitating membrane of birds and reptiles. Peritrophins, the integral PM proteins, are made of several domains. The major domain (peritrophin A-domain), is a cysteine-rich domain with chitinbinding properties having the consensus sequence: CX13–20CX5–6CX9–19CX10–14CX4–14C (where X is any amino acid except cysteine) that includes several conserved aromatic amino acids (see Section 4.5.4.5). Variations of this chitin-binding domain are peritrophin-B and peritrophin-C domains with consensus sequences: CX12–13CX20–21CX10CX12CX2CX8CX7–12C and CX8–9CX17–21CX10–11CX12–13CX11C, respectively. Another kind of domain occurring in peritrophins are proline/threonine-rich domains that are heavily glycosylated and similar to mucins. Peritrophins may have one (e.g., Cb-peritrophin-15 from Lucilia cuprina) to several (e.g., peritrophin-44 from

Biochemistry of Digestion

L. cuprina) chitin-binding domains or chitinbinding domains with small (e.g., Ag-AperI from Anopheles gambiae) or very large mucin-like domains (e.g., IIM from Trichoplusia ni) (Wang and Granados, 1997; Shen and Jacobs-Lorena, 1998; Tellam et al., 1999, 2003). The 3D structure of PM is thought to result from chitin fibrils being interlocked with the chitin-binding domains of peritrophins. Mucin-like domains of peritrophins are thought to face the ectoperitrophic and endoperitrophic sides of the PM. As these domains are highly hydrated they lubricate the surface of the PM, easing the movement of food inside the PM and of the ectoperitrophic fluid outside the PM. Furthermore, the glucan chains associated with peritrophin mucin-like domains may assure high proteinase resistance to PM (see Figure 9 in Schorderet et al., 1998; Figure 5 in Wang and Granados, 2001). The structure of peritrophins prompted Terra (2001) to develop a speculative model of the origin and evolution of the PM. According to this model, ancestral insects had their midgut cells covered with a mucus similar to that found in most animals. This gastrointestinal mucus, at least in vertebrates, is a gel-like substance composed of mucins (Allen, 1983; Forstner and Forstner, 1986). It was proposed that evolutionary processes led to the development of the PM from the gastrointestinal mucus. According to this hypothesis, the peritrophins, the major PM proteins, evolved from mucins by acquiring chitin– binding domains. The concomitant evolution of chitin secretion by midgut cells permitted the formation of the chitin–protein network characteristic of PM structure described above. Later on in evolution, some peritrophins lost their mucin-like domains. If the hypothesis that the PM is derived from the gastrointestinal mucus is correct, it should have originally been synthesized by midgut cells along the whole midgut and should have had the properties of the mucus. Later in evolution, insect species would have appeared with a chitin–protein network resulting in PM formation. Therefore, the formation of the PM by the whole midgut epithelium is the ancestral condition, whereas the restriction of PM production to midgut sections, or the lack of a PM and its replacement by the peritrophic gel, are derived conditions. PMs are classified into two types (Peters, 1992). Type I PM is found in cockroaches (Dictyoptera), grasshoppers (Orthoptera), beetles (Coleoptera), bees, wasps, and ants (Hymenoptera), moths and butterflies (Lepidoptera), and in hematophagous adult mosquitoes (Diptera). Type II PM occurs in larval

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and adult (except hematophagous ones) mosquitoes and flies (Diptera), and in a few adult Lepidoptera. Type I PM is formed either by the whole midgut epithelium, or by part of it (anterior or posterior regions). PM produced by the whole or anterior midgut epithelium envelops the food along the whole midgut. When PM is produced only by the posterior part, the anterior midgut epithelium is usually covered with a viscous material, the peritrophic gel, as observed in carabid beetles (Ferreira and Terra, 1989) and bees (Jimenez and Gilliam, 1990). This gel is also observed in the anteriorly placed midgut caeca of some insects and along the whole midgut of others (Terra, 2001). It has been shown with light microscopy, as well as transmission and scanning electron microscopy, that during the formation of type I PM chitincontaining fibrous material appears first at the tips of the microvilli of anterior midgut cells and then is rapidly included into a thin PM surrounding the food bolus (Harper and Hopkins, 1997). Chitin is synthesized outside the cells by a chitin synthase bound to microvillar membranes using precursors formed inside the cells (see Chapter 4.3). In agreement with this, antibodies raised against a translated product of a cDNA fragment coding chitin synthase immunolabeled only the apical ends of midgut microvilli (Zimoch and Merzendorfer, 2002). Chitin, after being self-organized into fibers, is interlocked by peritrophins. These are released by microapocrine secretion (Bolognesi et al., 2001). The formation of these PMs is frequently induced by the distension of the gut caused by food ingestion. Type II PM is secreted by a few rows of cells at the entrance of the midgut (cardia) and usually is found in insects irrespective of food ingestion. Peritrophins are secreted by exocytosis (Eisemann et al., 2001). Although a PM is found in most insects, it does not occur in Hemiptera and Thysanoptera, which have perimicrovillar membranes in their cells (see below). The other insects that apparently do not have a PM are adult Lepidoptera, Phthiraptera, Psocoptera, Zoraptera, Strepsiptera, Raphidioptera, Megaloptera, adult Siphonaptera, bruchid beetles, and some adult ants (Hymenoptera) (Peters, 1992). These insects may have a peritrophic gel instead of PM, one example being bruchid beetles (Terra, 2001), or may have had their PMs overlooked, because the insects were unfed. Another possibility is that minute hematophagous insects (like Siphonaptera and Phthiraptera) have lost their PM because the blood clot assures countercurrent flows (see Section 4.5.7.2.2.3) and their

200 Biochemistry of Digestion

small size makes easy the efficient diffusion of digestion products up to the midgut surface. The PM may have a large range of pore sizes: some small or very large and most of them in the middle range. The average pore sizes of PM may be determined by comparing molecular masses of enzymes restricted to the ectoperitrophic fluid (Figure 1) with those of enzymes present inside PM. This method of pore size estimation is probably the most accurate one since it reflects in vivo conditions. Pore sizes have also been determined by feeding insects with colloidal gold particles or fluorescent dextran molecules of known molecular masses and recording passage through the PM in vivo or using PM mounted as a sac and measuring diffusing rates. Determinations performed with these techniques by different authors (Zhuzhikov, 1964; Terra and Ferreira, 1983; Espinoza-Fuentes et al., 1984; Peters and Wiese, 1986; Santos and Terra, 1986; Wolfersberger et al., 1986; Miller and Lehane, 1990; Ferreira et al., 1994a) found pores in the range 7–9 nm for insects pertaining to different orders. Other authors described pores in the range 17–36 nm (Barbehenn and Martin, 1995; Edwards and Jacobs-Lorena, 2000). Pore sizes in the range 17–36 nm were obtained with fluorescent dextran molecules in conditions able to detect very small amounts of substances traversing the PM and, for this, they probably correspond to the large pores occurring at low frequency in PMs. Although these large pores are supposed to be of no importance regarding digestive events, they set the size limits that an infecting particle must have to successfully pass through the PM. As a consequences of its small pores, the PM hinders the free movement of molecules, dividing the midgut lumen into two compartments (Figure 1) with different molecules. The functions of this structure include those of the mucus (protection against food abrasion and invasion by microorganisms) and several roles associated with the compartmentalization of the midgut. These roles result in improvements in digestive physiology efficiency thereby leading to decreased digestive enzyme excretion, and restrict the production of the final products of digestion close to their transporters, thus facilitating absorption. These roles will be detailed below (see Section 4.5.7.2). In the light of current research, the distinction between PM and peritrophic gel may be more important than the traditional distinction between type I and type II PMs, in spite of the fact that the two types may have somewhat different structures (Tellam et al., 1999). Other major points needing clarification are how the chemical nature of

peritrophic gel and PM define their strength, elasticity and porosity and how these structures are self-assembled in the midgut lumen. 4.5.7.2. The Physiological Role of the Peritrophic Membrane

4.5.7.2.1. Protection against food abrasion and invasion by microorganisms As mentioned before, gut cells in most animals are covered with a gellike coating of mucus, which has been most thoroughly studied in mammals (Forstner and Forstner, 1986). In these animals, the mucus is supposed to lubricate the mucosa, protecting it from mechanical damage, and to trap bacteria and parasites. Since the insect midgut epithelium lacks a mucus coating, PM functions were supposed to be analogous to that of mucus. Thus, insects deprived of PM may have the midgut cells damaged by coarse food and may be liable to microorganism invasion in some reported cases (Peters, 1992; Tellam, 1996; Lehane, 1997). The PM as a barrier against invasion by microorganisms has particular relevance in insects that transmit viruses and parasites to human beings, as these microorganisms may have specific developmental phases in insect tissues (Tellam, 1996; Lehane, 1997). Microorganisms invade the insect midgut cells after disrupting the PM with the use of chitinase (Shahabuddin, 1995) or by using a proteinase such as enhancin that affects specifically the peritrophins (Peng et al., 1999; Ivanova et al., 2003). A barrier against microorganism invasion is probably less important for the majority of insects that feed on plants, as exemplified by observations carried out with the moth T. ni. Larvae of this insect deprived of PM by Calcofluor treatment show high mortality. Examination of dead larvae showed no signs of microbial infection or cell damage by Calcofluor, although these larvae were more susceptible to experimental infection (Wang and Granados, 2000). The results may be interpreted as Calcofluor killing larvae by affecting PM functions in digestion. In the same direction goes the observation that some plants respond to herbivorous insect attack by producing a unique 33 kDa cysteine proteinase with chitin-binding activity. This proteinase damages the PM, resulting in significant reduction in caterpillar growth caused by impaired nutrient utilization (Pechan et al., 2002). 4.5.7.2.2. Enhancing digestive efficiency 4.5.7.2.2.1. Overview The proposal of roles for the PM in digestion has benefited from studies on the organization of the digestive process. These studies (reviews: Terra, 1990; Terra and Ferreira,

Biochemistry of Digestion

1994, 2003) revealed that in most insects initial digestion occurs in the endoperitrophic space (Figure 1), intermediate digestion in the ectoperitrophic space, and final digestion at the surface of midgut cells. Such studies led to the formulation of the hypothesis of the endo–ectoperitrophic circulation

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of digestive enzymes. It was suggested that there is a recycling mechanism (Figure 11), where food flows inside the PM from the anterior midgut to the posterior, whereas in the ectoperitrophic space water flows from the posterior midgut to the caeca. When the polymeric food molecules become sufficiently

Figure 11 Diagrammatic representation of water fluxes (dotted arrows) and of the circulation of digestive enzymes (solid arrows) in putative insect ancestors that correspond to the major basic gut plans. In Neoptera ancestors (a), midgut digestive enzymes pass into the crop. Countercurrent fluxes depend on the secretion of fluid by the Malpighian tubules and its absorption by the caeca. Enzymes involved in initial, intermediate, and final digestion circulate freely among gut compartments. Holometabola ancestors (b) are similar except that secretion of fluid occurs in posterior ventriculus. The ancestors of hymenopteran and panorpoid (Lepidoptera and Diptera assemblage) insects (c) display countercurrent fluxes like Holometabola ancestors, midgut enzymes are not found in the crop, and only the enzymes involved in initial digestion pass through the peritrophic membrane. Enzymes involved in intermediate digestion are restricted to the ectoperitrophic space and those responsible for terminal digestion are immobilized at the surface of midgut cells. Cyclorrhapha ancestors (d) have a reduction in caeca, absorption of fluid in middle midgut, and anterior midgut playing a storage role. Lepidoptera ancestors (e) are similar to panorpoid ancestors, except that the anterior midgut replaces the caeca in fluid absorption. Hemiptera ancestors (f) have lost crop, caeca, and fluid-secreting regions. Fluid is absorbed in anterior midgut. (Reprinted with permission from Terra, W.R., Ferreira, C., 2003. Digestive system. In: Resh, V.H., Carde´, R.T. (Eds.), Encyclopedia of Insects. Academic Press, San Diego, CA, pp. 313–323; ß Elsevier.)

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small to pass through the PM (with the accompanying polymer hydrolases) the flow patterns result in the carriage towards the caeca or the anterior midgut where intermediate and final digestion occurs. Terra et al. (reviews: Terra and Ferreira, 1994; Terra, 2001) hypothesized that as a consequence of the compartmentalization of digestive events, there is an increase in the efficiency of digestion of polymeric food by allowing the removal of the oligomeric molecules from the endoperitrophic space, which is powered by the recycling mechanism associated with the midgut fluxes. Because oligomers may be substrates or inhibitors for some polymer hydrolases, their presence should decrease the rate of polymer degradation. Fast polymer degradation ensures that polymers are not excreted and hence increases their digestibility. Another possible consequence of compartmentalization is an increase in the efficiency of oligomeric food hydrolysis due to the transference of oligomeric molecules to the ectoperitrophic space and restriction of oligomer hydrolases to this compartment. In these conditions, oligomer hydrolysis occurs in the absence of probable partial inhibition (because of nonproductive binding) by polymer food and presumed nonspecific binding by nondispersed undigested food. This process should lead to the production of food monomers in the vicinity of midgut cell surface, causing an increase in the concentration of the final products of digestion close to their transporters, thus facilitating absorption. Experimental evidence supporting the adaptations for increasing digestive efficiency proposals are discussed in the following sections. 4.5.7.2.2.2. Prevention of nonspecific binding A model system was used to test the hypothesis that the PM prevents nonspecific binding of undigested material onto midgut cell surface (R. Bolognesi, W.R. Terra, and C. Ferreira, unpublished data). Aminopeptidase-containing microvillar membranes from Spodoptera frugiperda midgut cells were purified and assayed in the absence and presence of a concentration of midgut contents resembling in vivo conditions. The activity of the aminopeptidase in the presence of the midgut contents was about 50% of the activity in their absence, favoring the idea that undigested material in contact with microvillar enzymes negatively affects their activity. 4.5.7.2.2.3. Prevention of enzyme excretion This function was at first proposed based on results obtained with dipteran larvae (reviews: Terra, 1990; Terra and Ferreira, 1994). Both R. americana and Musca domestica present a decreasing trypsin gradient along midgut contents (putatively generated by

the recycling mechanism) and excreted less than 15% of midgut luminal trypsin after each gut emptying. When the larvae were fed a diet with excess protein, the trypsin gradient along midgut contents becomes less discernible and trypsin excretion increases to 40%. This is exactly what would be expected if the recycling mechanism existed and an increase in undigested dietary protein prevents trypsin from diffusing into the ectoperitrophic space and moving into anterior midgut by the countercurrent flux of fluid. Subsequently, dye experiments showed the existence of the appropriate fluid fluxes. More recently, experimental evidence that a recycling mechanism also occurs in Lepidoptera and Coleoptera was described. As predicted by the model, most trypsin activity is found in the lumen of Manduca sexta (Lepidoptera, type I PM) anterior midgut, whereas trypsin mRNA predominates in middle midgut (Peterson et al., 1994). Furthermore, immunocytochemical data showed the occurrence in the anterior midgut of significant amounts of a 41 kDa protein and a b-glycosidase secreted by the posterior midgut of M. sexta (Borhegyi et al., 1999) and middle midgut in Tenebrio molitor (Coleoptera, type I PM) (Ferreira et al., 2002), respectively. Finally, the decreasing trypsin and chymotrysin gradient along S. frugiperda midgut contents disappeared in Calcofluor-treated larvae lacking a peritrophic membrane (Bolognesi et al., 2001) and the excretory rate increased from 0.1% to 0.9% of midgut contents at each gut emptying (R. Bolognesi, W. R. Terra, and C. Ferreira, unpublished data). 4.5.7.2.2.4. Increase in the efficiency of digestion of polymeric food A model system was used to test this hypothesis (R. Bolognesi, W.R. Terra, and C. Ferreira, unpublished data). Midgut contents from S. frugiperda larvae were placed into dialysis bags suspended in stirred and unstirred media. Trypsin activities in stirred and unstirred bags were 210% and 160%, respectively, over the activities of similar samples maintained in a test tube. The results suggested that the diffusion of products from the trypsin reaction media favors enzyme action and that stirring (an in vitro model of the ectoperitrophic countercurrent flux) enhances the effect. 4.5.7.2.2.5. Increase in the efficiency of oligomeric food hydrolysis This putative function is supported by the experiments of Bolognesi et al. (R. Bolognesi, W.R. Terra, and C. Ferreira, unpublished data). They collected ectoperitrophic fluid from the large midgut caeca of R. americana. Aminopeptidase A, N-acetylglucosaminidase, and carboxypeptidase A are enzymes restricted to the

Biochemistry of Digestion

ectoperitrophic space. When those enzymes were put in the presence of PM contents their activities decreased in relation to controls as follows: aminopeptidase A, 46%; N-acetylglucosaminidase, 56%; carboxypeptidase A, 92%. These decreases in activity probably result from oligomer hydrolase competitive inhibition by luminal polymers. 4.5.7.2.2.6. Restriction of food monomer production at cell surface This is a consequence of restricting oligomer hydrolases to the ectoperitrophic space (see Section 4.5.7.2.2.5) and causes an increase in the concentration of the final products of digestion close to the carriers responsible for their absorption. A model system should be developed to test this hypothesis. 4.5.7.2.2.7. Enzyme immobilization Midgut luminal enzymes, in addition to occurring in the endoperitrophic and ectoperitrophic spaces, may be associated with the PM. For example, results obtained with S. frugiperda larvae showed that PM may contain up to 13% and 18% of the midgut luminal activity of amylase and trypsin, respectively (Ferreira et al., 1994b). Hence, enzyme immobilization may play a role in digestion, although a minor one. The attachment mechanism of enzymes in PM is not well known. Nevertheless, there is evidence, at least in S. frugiperda, that trypsin, amylase, and microvillar enzymes are incorporated into the jelly-like substance associated with PM when the enzymes, still bound to membranes, are released from midgut cells by a microaprocrine process (Jorda˜ o et al., 1999; Bolognesi et al., 2001). 4.5.7.2.2.8. Toxin binding Although potentially toxic dietary tannins are attached to and excreted with Schistocerca gregaria PM (Bernays and Chamberlain, 1980), toxin binding by the PM seems to be a less widespread phenomenon than previously suggested. Thus, tannins in M. sexta (Barbehenn and Martin, 1998) and lipophilic and amphiphilic noxious substances in Melanoplus sanguinipes (Barbehenn, 1999) are maintained in the endoperitrophic space because they form high molecular weight complexes, not because of PM binding. 4.5.7.2.2.9. Peritrophic membrane functions and insect phylogeny Current data detailed below suggest that PMs of all insects have functions (see Sections 4.5.7.2.1, 4.5.7.2.2.2–4.5.7.2.2.4), whereas functions (see Sections 4.5.7.2.2.5 and 4.5.7.2.2.6) are demonstrable only in PMs of Panorpodea (the taxon that includes Diptera and

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Lepidoptera) and of the hymenopteran sawflies. PM function (see Section 4.5.7.2.2.7) may occur in all insects but this needs further confirmation. Function (see Section 4.5.7.2.2.8), although it may be important for some insects, should be viewed as opportunistic. In other words, the PM probably evolved from a protective role (see Section 4.5.7.2.1) to more sophisticated functions (see Sections 4.5.7.2.2.2–4.5.7.2.2.7) under selective pressures and, due to the chemical properties of their constituents, the PM also developed the ability to bind different compounds including toxins.

4.5.8. Organization of the Digestive Process 4.5.8.1. Evolutionary Trends of Insect Digestive Systems

After studying the spatial organization of the digestive events in insects of different taxa and diets, it was realized that insects may be grouped relative to their digestive physiology, assuming they have common ancestors.Those putative ancestors correspond to basic gut plans from which groups of insects may have evolved by adapting to different diets (Terra and Ferreira, 1994, 2003). The basic plan of digestive physiology for most winged insects (Neoptera ancestors) is summarized in Figure 11. In these ancestors, the major part of digestion is carried out in the crop by digestive enzymes propelled by antiperistalsis forward from the midgut. Saliva plays a variable role in carbohydrate digestion. After a while following ingestion, the crop contracts transferring digestive enzymes and partly digested food into the ventriculus. The anterior ventriculus is acid and has high carbohydrase activity, whereas the posterior ventriculus is alkaline and has high proteinase activity. This differentiation along the midgut may be an adaptation to instability of ancestral carbohydrases in the presence of proteinases. The food bolus moves backward in the midgut of the insect by peristalsis. As soon as the polymeric food molecules are digested to become sufficiently small to pass through the peritrophic membrane, they diffuse with the digestive enzymes into the ectoperitrophic space (Figure 1). The enzymes and nutrients are then displaced toward the caeca with a countercurrent flux caused by secretion of fluid at the Malpighian tubules and its absorption back by cells at the caeca (Figure 11), where final digestion is completed and nutrient absorption occurs. When the insect starts a new meal, the caeca contents are moved into the crop. As a consequence of the countercurrent flux,

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digestive enzymes occur as a decreasing gradient in the midgut and their excretion is lowered. The Neoptera basic plan gave origin to that of the Polyneoptera orders, which include Blattodea, Isoptera, and Orthoptera, and evolved to the basic plans of Paraneoptera and Holometabola. The characteristics of the Paraneoptera ancestors cannot be inferred because midgut function data are available only for Hemiptera. The basic gut plan of the Holometabola (Figure 11b) (which include Coleoptera, Megaloptera, Hymenoptera, Diptera, and Lepidoptera) is similar to that of Neoptera, except that fluid secretion occurs by the posterior ventriculus, instead of by the Malpighian tubules. Because the posterior midgut fluid does not contain wastes, as is the case for Malpighian tubular fluid, the accumulation of wastes in caeca is decreased. Caeca loss probably further decreases the accumulation of noxious substances in the midgut, which would be more serious in insects that have high relative food consumption rates, as is common among Holometabola. The basic plan of Coleoptera did not evolve dramatically from the holometabolan ancestor, whereas the basic plan of Hymenoptera, Diptera, and Lepidoptera ancestor (hymenopteran–panorpoid ancestor, Figure 11c) presents important differences. Thus, hymenopteran–panorpoid ancestors have countercurrent fluxes like holometabolan ancestors, but differ from these in the lack of crop digestion, midgut differentiation in luminal pH, and in which compartment is responsible for each phase of digestion. In holometabolan ancestors, all phases of digestion occur in the endoperitrophic space (Figure 1), whereas in hymenopteran–panorpoid ancestors only initial digestion occurs in that region. In the latter ancestors, intermediate digestion is carried out by free enzymes in the ectoperitrophic space and final digestion occurs at the midgut cell surface by immobilized enzymes. The free digestive enzymes do not pass through the PM because they are larger than the PM’s pores. As a consequence of the compartmentalization of digestive events in hymenopteran and panorpoid insects, there is an increase in the efficiency of digestion of polymeric food as discussed before. The evolution of insect digestive systems summarized above and in Figure 11 was proposed, as discussed before, from studies carried out in 12 species pertaining to four insect orders. To give further support for the hypothesis that the characteristics of gut function and morphology depend more on phylogeny than on diet, another approach was used. A total of 29 gut morphology and digestive physiology characteristics (e.g., luminal pH, ratio of gut

Figure 12 Cladogram of representative insects based on 29 gut morphology and digestive physiology characteristics. Insects: 1, Trichosia pubescens; 2, Rhynchosciara americana; 3, Musca domestica; 4, Anopheles spp.; 5, Rhodnius prolixus; 6, Dysdercus peruvianus; 7, Acyrthosiphon pisum; 8, Erinnyis ello; 9, Spodoptera frugiperda; 10, Themos malaisei; 11, Camponotus rufipes; 12, Scaptotrigona bipunctata; 13, Bracon hebetor; 14, Tenebrio molitor; 15, Migdolus fryanus; 16, Sphenophorus levis; 17, Cyrtomon solana; 18, Dermestes maculatus; 19, Pyrearinus termitilluminans; 20, Pheropsophus aequinoctialis; 21, Corydalus sp.; 22, Abracris flavolineata; 23, Periplaneta americana. (Courtesy of A.B. Dias.)

section volumes, type of peritrophic membrane, presence of special gut cells, distribution of digestive enzymes along the gut, major proteinase) were identified in 23 species from eight different insect orders. Making use of these characteristics, a cladogram was constructed putting together the data from studied species (Figure 12). The data confirmed that the morphological and functional traits associated with the digestive system are more dependent on taxon than on dietary habits of the different insects (Dias, Vanin, Marques, and Terra, unpublished data). There are two insect species that do not apparently fit the model: Anopheles spp. and Themos malaisei. Anopheles spp. is an adult, whereas the other Diptera is larval. Themos malaisei is an unexpected finding that will be discussed below (see Section 4.5.8.2.8). 4.5.8.2. Digestion in the Major Insect Orders

4.5.8.2.1. Initial comments Applebaum (1985) in his review for the first edition of this series divided the insects according to their diet when describing their digestive processes. Recent data (see Section 4.5.8.1) support the view that functional digestive traits of insects are linked with their phylogenetic

Biochemistry of Digestion

position. The organization of the digestive process in the different insect orders have been reviewed several times (Terra, 1988, 1990; Terra and Ferreira, 1994, 2003). The following section is therefore an abridged version of those texts, highlighting new findings and trying to identify points that deserve more research, especially in relation to molecular aspects. Only key references before 1994 are cited and the reader should find more references in the above-mentioned reviews. 4.5.8.2.2. Blattodea Cockroaches are usually omnivorous. It is thought that digestion in cockroaches occurs as described for the Neoptera ancestor (Figure 11a), except that part of the final digestion of proteins occurs on the surface of midgut cells (Terra and Ferreira, 1994). This was confirmed by the finding in P. americana that most trypsin, maltase, and amylase are found in the crop, whereas aminopeptidase predominates in the microvillar membranes of posterior midgut. There is a decreasing gradient of trypsin, maltase, and amylase along the midgut contents and less than 5% of trypsin and maltase (amylase, 27%) are excreted during each midgut emptying. This suggests the existence of midgut digestive enzyme recycling, with amylase excretion increased probably due to excess dietary starch. The recycling mechanism is thought to be powered by water fluxes as in the Neoptera ancestor, although there are no data supporting this. Major digestive proteinases are trypsin and chymotrypsin (Dias and Terra, unpublished data). The differentiation of pH along the midgut (acid anterior midgut and alkaline posterior midgut) is not conserved among some cockroaches like P. americana, but it was maintained in others exemplified by the blaberid Nauphoeta cinerea (Elpidina et al., 2001a). The organization of digestion in this cockroach seems similar to that in P. americana, although data on enzyme excretion are lacking. At least blaberoid cockroaches possess proteinase inhibitory proteins active in the anterior midgut. These inhibitors are thought to be a primitive device to decrease the proteolytic inactivation of the animal’s own carbohydrases, which are thus expected to be more active in the anterior midgut. The digestive carbohydrases from more evolved insects are stable in the presence of their own proteinases (Terra, 1988). Recently several proteinase inhibitors have been partially purified from N. cinerea (Elpidina et al., 2001b). Another difference between cockroaches and the Neoptera ancestor is the enlargement of hindgut structures, noted mainly in wood-feeding cockroaches. These hindgut structures harbor

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bacteria producing acetate and butyrate from ingested wood or other cellulose-containing materials. Acetate and butyrate are absorbed by the hindgut of all cockroaches, but this is more remarkable with woodroaches (Terra and Ferreira, 1994). Cellulose digestion may be partly accomplished by bacteria in the hindgut of P. americana or protozoa in Cryptocercus punctulatus (Bignell, 1981). Nevertheless, now it is clear that P. americana saliva contains two cellulases and three laminarinases that may open plant cells and lyze fungal cells (Genta et al., 2003). This agrees with the omnivorous detritus feeding habit of the insect. The woodroach Panestria cribrata also has its own cellulase (Scrivener et al., 1989). 4.5.8.2.3. Isoptera Termites may be seen as insects derived from, and more adapted than, woodroaches in dealing with refractory material as wood and humus. Associated with this specialization, they lost the crop and midgut caeca, and enlarged their hindgut structures. Both lower and higher termites digest cellulose with their own cellulase, despite the occurrence of cellulose-producing protozoa in the paunch, an enlarged region of the anterior hindgut in lower termites. The products of cellulose digestion pass from the midgut into the hindgut, where they are converted into acetate and butyrate by hindgut bacteria as in woodroaches. Symbiotic bacteria are also responsible for nitrogen fixation in the hindgut, resulting in bacterial protein. This is incorporated into the termite body mass after being expelled in feces by one individual and being ingested and digested by another. This explains the ability of termites to develop successfully in diets very poor in protein. Both lower and higher feeding termites seem to have an endo–ectoperitrophic circulation of digestive enzymes (Terra and Ferreira, 1994; Nakashima et al., 2002; see also Section 4.5.4.3.1). 4.5.8.2.4. Orthoptera Grasshoppers feed mainly on grasses and their digestive physiology has clearly evolved from the Neoptera ancestor. Carbohydrate digestion occurs mainly in the crop, under the action of midgut enzymes, whereas protein digestion and final carbohydrate digestion take place at the anterior midgut caeca. The abundant saliva (devoid of significant enzymes) produced by grasshoppers saturate the absorbing sites in the midgut caeca, thus hindering the countercurrent flux of fluid. This probably avoids excessive accumulation of noxious wastes in the caeca, coming from Malpighian tubule secretion, and makes possible the high relative food consumption observed among locusts

206 Biochemistry of Digestion

in their migratory phases. Starving grasshoppers present midgut countercurrent fluxes. Cellulase found in some grasshoppers is believed to facilitate the access of digestive enzymes to the plant cells ingested by the insects by degrading the cellulose framework of cell walls (Dow, 1986; Terra and Ferreira, 1994; Marana et al., 1997). 4.5.8.2.5. Hemiptera The Hemiptera comprise insects of the major suborders Auchenorrhyncha (cicadas, spittlebugs, leafhoppers, and planthoppers) and Sternorrhyncha (aphids and white flies) that feed almost exclusively on plant sap, and Heteroptera (e.g., assassin bugs, plant bugs, stink bugs, and lygaeid bugs) that are adapted to different diets. The ancestor of the entire order Hemiptera is supposed to have been a sap-sucker similar to present day Auchenorrhyncha. Sap-sucking Hemiptera may suck phloem or xylem sap. These food sources have very low contents of proteins (with the exception of few phloem saps; see below) and carbohydrate polymers and are relatively poor in free essential amino acids. In contrast to xylem sap, phloem sap is very rich in sucrose (Terra, 1990). Thus, except for dimer (sucrose) hydrolysis, no food digestion is usually necessary in sap-suckers. Upon adapting to dilute phloem and/or xylem sap, hemipteran ancestors would lose the enzymes involved in initial and intermediate digestion and lose the peritrophic membrane (Figure 11f). These changes are associated with the lack of luminal digestion. The major problem facing a sap-sucking insect (specially on dilute phloem or xylem sap) is to absorb nutrients, such as essential amino acids, that are present in very low concentration in sap. Whichever mechanism is used, xylem feeders may absorb as much as 99% of dietary amino acids and carbohydrate (Andersen et al., 1989). Amino acids may be absorbed according to a hypothesized mechanism that depends on perimicrovillar membranes, which are membranes ensheathing the midgut microvilli with a dead end (Figure 13). A role in midgut amino acid absorption depends on the presence of amino acid–Kþ symports on the surface of the perimicrovillar membranes and of amino acid carriers and potassium pumps on the microvillar membranes. Although amino acid carriers have been found in the microvillar membranes of several insects (Wolfersberger, 2000), no attempts have been made to study the other postulated proteins. Thus, in spite of the model provided an explanation for the occurrence of these peculiar cell structures in Hemiptera, it is supported only by: (1) evidence that amino acids are absorbed with potassium ions in

Dysdercus peruvianus (Silva and Terra, 1994); (2) occurrence of particles studying the cytoplasmic face of the midgut microvillar membranes of D. peruvianus. These might be ion pumps responsible for the putative potassium ion transport, like similar structures in several epithelia (Silva et al., 1995). Another problem that deserves more attention regarding perimicrovillar membranes is their origin. Immunolocalization of the perimicrovillar enzyme marker, a-glucosidase, suggests that these membranes are formed when double membrane vesicles fuse their outer membranes with the microvillar membranes and their inner membranes with the perimicrovillar membranes. A double membrane Golgi cisterna (on budding) forms the double membrane vesicles (Silva et al., 1995). Organic compounds in xylem sap need to be concentrated before they can be absorbed by the perimicrovillar system. This occurs in the filter chamber of Cicadoidea and Cercopoidea, which concentrates xylem sap tenfold. The filter chamber consists of a thin-walled, dilated anterior midgut in close contact with the posterior midgut and the proximal ends of the Malpighian tubules. This arrangement enables water to pass directly from the anterior midgut to the Malpighian tubules, concentrating food in midgut and eliminating excess water. The high permeability of the filter chamber membrane to water results from the occurrence of specific channels formed by proteins named aquaporins. These were characterized as membrane proteins with 15–26 kDa and were immunolocalized in the filter chamber of several xylem sap feeders (Le Cahe´ rec et al., 1997). Sternorrhyncha, as exemplified by aphids, may suck more or less continuously phloem sap of sucrose concentration up to 1.0 M and osmolarity up to three times that of the insect hemolymph. This results in a considerable hydrostatic pressure caused by the tendency of water to move from the hemolymph into midgut lumen. To withstand these high hydrostatic pressures, aphids have developed several adaptations. Midgut stretching resistance is helped by the existence of links between apical lamellae (replacing usual midgut cell microvilli) that become less conspicuous along the midgut. As a consequence of the links between the lamellae, the perimicrovillar membranes could no longer exist and were replaced by membranes seen associated with the tips of the lamellae, the modified perimicrovillar membranes (Ponsen, 1991; Cristofoletti et al., 2003). A modified perimicrovillar membrane-associated a-glucosidase frees fructose from sucrose without increasing the osmolarity by promoting transglycosylations. As the fructose is quickly absorbed, the osmolarity decreases, resulting in a

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Figure 13 Model for the structure and physiological role of the microvillar border of midgut cells from Hemiptera. The left figure is a diagrammatic representation of a typical Hemiptera midgut cell and the right figure details its apex. The microvillar membrane (MM) is ensheathed by the perimicrovillar membrane (PMM), which extends toward the luminal compartment with a dead end. The microvillar and perimicrovillar membranes delimit a closed compartment, i.e., the perimicrovillar space (PMS). The microvillar membrane is rich and the perimicrovillar membrane is poor in integral proteins (IP). Microvillar membranes actively transport potassium ions (the most important ion in sap) from PMS into the midgut cells, generating a concentration gradient between the gut luminal sap and the PMS. This concentration gradient may be a driving force for the active absorption of organic compounds (amino acids, aa, for example) by appropriate carriers present in the PMM. Organic compounds, once in the PMS, may diffuse up to specific carriers on the microvillar surface. This movement is probably enhanced by a transfer of water from midgut lumen to midgut cells, following (as solvation water) the transmembrane transport of compounds and ions by the putative carriers. (Reprinted with permission from Terra, W.R., Ferreira, C., 1994. Insect digestive enzymes: properties, compartmentalization and function. Comp. Biochem. Physiol. B 109, 1–62; ß Elsevier.)

honeydew isoosmotic with hemolymph (Ashford et al., 2000; Cristofoletti et al., 2003). Another interesting adaptation is observed in whiteflies, where a trehalulose synthase forms trehalulose from sucrose, thus making available less substrate for an a-glucosidase that otherwise would increase the osmolarity of ingested fluid on hydrolyzing sucrose (Salvucci, 2003). A cathepsin L (see Section 4.5.5.3) bound to the modified perimicrovillar membranes of Acyrthosiphon pisum (Cristofoletti et al., 2003) may explain the capacity of some phloem sap feeders to rely on protein found in some phloem saps (Salvucci et al., 1998) and the failure of other authors to find an active proteinase in sap feeders. They worked with homogenate supernatants or supernatants of Triton

X-100-treated samples, under which conditions the cathepsin L would remain in the pellet. Amino acid absorption in A. pisum midguts is influenced by the presence of the bacteria Buchnera in the mycetocytes of the mycetomes occurring in the aphid hemocoel (Prosser et al., 1992). The molecular mechanisms underlying this phenomenon are not known, in spite of the fact that there is strong evidence showing that Buchnera uses the nonessential amino acids absorbed by the host in the synthesis of essential amino acids (Prosser and Douglas, 1992; Shigenobu et al., 2000). It is likely that amino acid absorption through apical lamellar carriers depends on the amino acid concentration gradient between midgut lumen and hemolymph, whereas hemolymph titers vary widely according

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p0925

to Buchnera metabolic activity (Liadouze et al., 1995). The evolution of Heteroptera was associated with regaining the ability to digest polymers. Because the appropriate digestive enzymes were lost, they instead used enzymes derived from lysosomes. Lysosomes are cell organelles involved in intracellular digestion carried out by special proteinases referred to as cathepsins. Compartmentalization of digestion was maintained by the perimicrovillar membranes, as a substitute for the absent peritrophic membrane. Digestion in the two major Heteroptera taxa, Cimicomorpha (exemplified by the blood-feeder Rhodnius prolixus), and Pentatomorpha (exemplified by the seed-sucker Dysdercus peruvianus), are similar. The dilated anterior midgut stores food and absorbs water and, at least in D. peruvianus, also absorbs glucose. Digestion of proteins and absorption of amino acids occurs in the posterior ventriculus. Most protein digestion occurs in lumen with the aid of a cysteine proteinase, and ends in the perimicrovillar space under the action of aminopeptidases and dipeptidases (Terra and Ferreira, 1994). Symbiont bacteria may occur in blood-feeders putatively to provide vitamins (see Section 4.5.2.5). At least in R. prolixus, the neuroendocrine system has factors important for maintaining the ultrastructural organization of the midgut epithelial cells (Gonzales et al., 1998). 4.5.8.2.6. Megaloptera Megaloptera include alderflies and dobsonflies and are often considered to be the most primitive group of insects with complete metamorphosis. All their larvae are aquatic predators feeding on invertebrates (Theischinger, 1991). Megaloptera ancestors are like Holometabola ancestors except that for the anterior midgut caeca, which were lost and replaced in function by the anterior midgut. Thus, in Corydalus sp. larvae, most digestion occurs in the crop under the action of soluble amylase, maltase, aminopeptidase, and trypsin (major proteinase). Digestive enzyme recycling should occur, as less than 10% of midgut amylase, maltase, and aminopeptidase are lost at each midgut emptying. The higher excretory rate of trypsin (27%) probably results from excess dietary protein (Dias and Terra, unpublished data). 4.5.8.2.7. Coleoptera Coleoptera ancestors are like Megaloptera ones. Nevertheless, there are evolutionary trends leading to a great reduction or loss of the crop and, as in the panorpoid orders, occurrence of at least final digestion of proteins at the surface of midgut cells. Thus, in predatory Carabidae most of the digestive phases occur in the crop by

means of midgut enzymes, whereas in predatory larvae of Elateridae initial digestion occurs extraorally by the action of enzymes regurgitated onto their prey. The preliquified material is then ingested by the larvae and its digestion is finished at the surface of midgut cells (Terra and Ferreira, 1994). The entire digestive process occurs in the dermestid larval endoperitrophic space that is limited by a peritrophic gel in anterior midgut and a peritrophic membrane in posterior midgut. There is a decreasing gradient along the midgut of amylase, maltase, trypsin (major proteinase), and aminopeptidase suggesting the occurrence of digestive enzyme recycling (Terra and Ferreira, 1994; Caldeira, Dias, Terra, and Ribeiro, unpublished data). Like dermestid beetles, the larvae of Migdolus fryanus (Cerambycidae) and Sphenophorus levis (Curculionidae) have a peritrophic gel and a peritrophic membrane in the anterior and posterior midgut, respectively, and a decreasing gradient of amylase, maltase, and proteinase along the midgut. In contrast to dermestids, aminopeptidase is a microvillar enzyme in both insects (Dias and Terra, unpublished data). These data do not confirm the earlier suggestion (Terra and Ferreira, 1994) that the final digestion of all nutrients occurs on the surface of midgut cells of Curculionidae. Tenebrionid larvae also have aminopeptidase as a microvillar enzyme and the distribution of enzymes in gut regions of adults is similar to that in the larvae (Terra and Ferreira, 1994). This suggests that the overall pattern of digestion in larvae and adults of Coleoptera is similar, despite the fact that (in contrast to adults) beetle larvae usually lack a crop. Insects of the series Cucujiformia (which includes Tenebrionidae, Chrysomelidae, Bruchidae, and Curculionidae) have cysteine proteinases (see Section 4.5.5.3) in addition to (or in place of) serine proteinases as digestive enzymes, suggesting that the ancestors of the whole taxon were insects adapted to feed on seeds rich in serine proteinase inhibitors. The occurrence of trypsin as the major proteinase in M. fryanus (Dias and Terra, unpublished data) confirmed the preliminary work (Murdock et al., 1987) according to which cerambycid larvae reacquired serine proteinases. Scarabaeidae and several related families are relatively isolated in the series Elateriformia and evolved considerably from the Coleoptera ancestor. Scarabid larvae, exemplified by dung beetles, usually feed on cellulose materials undergoing degradation by a fungus-rich flora. Digestion occurs in the midgut, which has three rows of caeca, with a ventral groove between the middle and posterior row. The alkalinity of gut contents increase to almost

Biochemistry of Digestion

pH 12 along the midgut ventral groove. This high pH probably enhances cellulose digestion, which occurs mainly in the hindgut fermentation chamber, through the probable action of bacterial cell-bound enzymes. The final product of cellulose degradation is mainly acetic acid, which is absorbed through the hindgut wall. There is controversy as to whether scarabid larvae ingest feces to obtain nitrogen compounds, as described above for termites (Terra and Ferreira, 1994; Biggs and McGregor, 1996). 4.5.8.2.8. Hymenoptera The organization of the digestive process is variable among hymenopterans and to understand its peculiarities it is necessary to review briefly their evolution. The hymenopteran basal lineages are phytophagous as larvae, feeding both ecto- and endophytically and include several superfamilies like Xyeloidea and Tenthredinoidea, all known as sawflies. Close to these are the Siricoidea (wood wasps) that are adapted to ingest fungus-infected wood. Wood wasp-like ancestors gave rise to the Apocrita (wasp-waisted Hymenoptera) that are parasitoids of insects. They use their ovipositor to injure or kill their host which represents a single meal for their complete development. A taxon sister of Ichneumonoidea in Apocrita gave rise to Aculeata (bees, ants, and wasps with thin waist) (Quicke, 2003). The digestive systems of Hymenoptera ancestors are like the panorpoid ancestors (Figure 11c). However, there are evolutionary trends leading to the loss of midgut caeca (replaced in function by the anterior midgut) and changes in midgut enzyme compartmentalization. These trends appear to be associated with the development of parasitoid habits and were maintained in Aculeata, as described below. The sawfly T. malaisei (Tenthredinoidea: Argidae) larva has a midgut with a ring of anterior caeca that forms a U at the ventral side. Luminal pH is above 9.5 in the first two-thirds of the midgut. Trypsin (major proteinase) and amylase have a decreasing activity along the endoperitrophic space, suggesting enzyme recycling. Maltase predominates in the anterior midgut tissue as a soluble glycocalyx-associated enzyme, whereas aminopeptidase is a microvillar enzyme in posterior midgut (Dias, Ribeiro, and Terra, unpublished data). These characteristics (except the presence of caeca) are similar to those of lepidopteran larvae (see Section 4.5.8.2.10) and explain the fact that this insect is close to the lepidopterans in Figure 12. Otherwise, Aculeata with their less sophisticated midgut (see below) branches closer to coleopterans (Figure 12). Wood wasp larvae of the genus Sirex are believed to be able to digest and assimilate wood constituents

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by acquiring cellulase and xylanase, and possibly other enzymes, from fungi present in wood on which they feed (Martin, 1987). The larvae of Apocrita present a midgut which is closed at its rear end, and which remains unconnected with the hindgut until the time of pupation. It is probable that this condition evolved as an adaptation of endoparasitoid Apocrita ancestors to avoid the release of toxic compounds into the host in which they lived (Terra, 1988). In larval bees, most digestion occurs in the endoperitrophic space. Countercurrent fluxes seem to occur but there is no midgut luminal pH gradient. Adult bees ingest nectar and pollen. Sucrose from nectar is hydrolyzed in the crop by the action of a sucrase from the hypopharyngeal glands. After ingestion, pollen grains extrude their protoplasm in the ventriculus, where digestion occurs. As in larvae there is also evidence of an endo–ectoperitrophic circulation of digestive enzymes (Jimenez and Gilliam, 1990; Terra and Ferreira, 1994). Although many authors favor the view that pollen grains are digested in bees after their extrusion by osmotic shock, this subject is controversial not only in bees but also among pollen-feeder beetles (Human and Nicholson, 2003). Worker ants feed on nectar, honeydew, plant sap, or on partly digested food regurgitated by their larva. Thus, they frequently were said to lack digestive enzymes or display only those enzymes associated with intermediate and (or) final digestion (Terra and Ferreira, 1994). Although this seems true for leaf-cutting ants that appear to rely only on monosaccharides, produced by fungal enzymes acting on plant polysaccharides (Silva et al., 2003), this is not widespread. Thus, adult Camponotus rufipes (Formicinae) have soluble amylase, trypsin (major proteinase), maltase, and aminopeptidase enclosed in a type I PM in their midguts. As only 14% of amylase and less than 7% of the other digestive enzymes are excreted during the midgut emptying, these insects may have a digestive enzyme recycling mechanism (Dias and Terra, unpublished data). 4.5.8.2.9. Diptera The Diptera evolved along two major lines: an assemblage (early Nematocera) of suborders corresponding to the mosquitoes, including the basal Diptera, and the suborder Brachycera that includes the most evolved flies (Cyclorrhapha). The Diptera ancestor is similar to the panorpoid ancestor (Figure 11c) in having the enzymes involved in intermediate digestion free in the ectoperitrophic fluid (mainly in the large caeca), whereas the enzymes of terminal digestion are

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membrane bound at the midgut cell microvilli (Terra and Ferreira, 1994). Although these characteristics are observed in most nonbrachyceran larvae, the more evolved of these larvae may show reduction in size of midgut caeca (e.g., Culicidae). Nonhematophagous adults store liquid food (nectar or decay products) in their crops. Digestion occurs in their midgut as in larvae. Nectar ingested by mosquitoes (males and females) is stored in the crop, and is digested and absorbed at the anterior midgut. Blood, which is ingested only by females, passes to the posterior midgut, where it is digested and absorbed (Billingsley, 1990; Terra and Ferreira, 1994). Adult Aedes aegypti midgut surface is covered in a large proportion by tubular bilayers with diameter fourfold smaller than microvilli. They fuse and branch forming bundles that seem to originate in the intercellular crypts and seem to be fused with the microvillar surface (Ziegler et al., 2000). These structures are not related with the perimicrovillar membranes of Hemiptera. The latter envelop the microvilli before extending into the lumen in structures that may resemble the tubular membrane bilayers of A. aegypti (see Section 4.5.8.2.5 and Figure 13). The puzzling structures of A. aegypti should be further studied to discover their relationships with digestion. The Cyclorrhapha ancestor (Figure 11d) evolved dramatically from the panorpoid ancestor (Figure 11c), apparently as a result of adaptations to a diet consisting mainly of bacteria. Digestive events in Cyclorrhapha larvae are exemplified by larvae of the housefly Musca domestica. These ingest food rich in bacteria. In the anterior midgut there is a decrease in the starch content of the food bolus, facilitating bacterial death. The bolus now passes into the middle midgut where bacteria are killed by the combined action of low pH, a special lysozyme (see Section 4.5.4.5) and an aspartic proteinase (see Section 4.5.5.4). Finally, the material released by bacteria is digested in the posterior midgut, as is observed in the whole midgut of insects of other taxa. Countercurrent fluxes occur in the posterior midgut powered by secretion of fluid in the distal part of the posterior midgut and its absorption back in middle midgut. The middle midgut has specialized cells for buffering the luminal contents in the acidic zone (Figure 4), in addition to those functioning in fluid absorption. Cyclorrhaphan adults, except for a few blood-suckers, feed mainly on liquids associated with decaying material (rich in bacteria) in a way similar to housefly M. domestica adults. These salivate (or regurgitate their crop contents) onto their food. After the dispersed material is ingested, starch digestion is accomplished

primarily in the crop by the action of salivary amylase. Digestion is followed in the midgut, essentially as described for larvae (Terra and Ferreira, 1994). The stable fly Stomoxys calcitrans stores and concentrates the blood meal in the anterior midgut and gradually passes it to the posterior midgut, where digestion takes place, resembling what occurs in larvae. These adults lack the characteristic cyclorrhaphan middle midgut and the associated luminal low pH. Stable flies occasionally take nectar (Jorda˜ o et al., 1996a). 4.5.8.2.10. Lepidoptera Lepidopteran ancestors (Figure 11e) differ from panorpoid ancestors because they lack midgut caeca, have all their digestive enzymes (except those of initial digestion) immobilized at the midgut cell surface, and present long-neck goblet cells and stalked goblet-cells in the anterior and posterior larval midgut regions, respectively. Goblet cells excrete Kþ ions that are absorbed from leaves ingested by larvae. Goblet cells also seem to assist anterior columnar cells in water absorption and posterior columnar cells in water secretion (Terra and Ferreira, 1994; Ortego et al., 1996). Although most lepidopteran larvae have a common pattern of digestion, species that feed on unique diets generally display some adaptations. Tineola bisselliella (Tineidae) larvae feed on wool and display a highly reducing midgut for cleaving the disulfide bonds in keratin to facilitate proteolytic hydrolysis of this otherwise insoluble protein (Terra and Ferreira, 1994). Similar results were obtained with Hofmannophila pseudospretella (Christeller, 1996). Wax moths (Galleria mellonella) infest beehives and digest and absorb wax. The participation of symbiotic bacteria in this process is controversial. Another adaptation has apparently occurred in lepidopteran adults which feed solely on nectar. Digestion of nectar only requires the action of an a-glucosidase (or a b-fructosidase) to hydrolyze sucrose, the major component present. Many nectar-feeding lepidopteran adults have amylase in salivary glands and several glycosidases and peptidases in the midgut (Terra and Ferreira, 1994). Woods and Kingsolver (1999) developed a chemical reactor model of the caterpillar midgut and used the model as a framework for generating hypotheses about the relationship between feeding responses to variable dietary protein and the physical and biochemical events in the midgut and body. They concluded that absorption (or postabsorptive processes) is limiting in a caterpillar maintained in artificial diets. Caterpillars eating leaves may not have

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the same limiting step and this deserves a similar detailed study. Another interesting study would be the development of a model for the beetle midgut. This would determine whether beetles have digestion as the limiting step or consumption to compensate for their less sophisticated midguts.

4.5.9. Digestive Enzyme Secretion Mechanisms and Control Digestive enzyme secretory mechanisms and control probably are the least understood areas in insect digestion. Studies of the secretory mechanisms have only described major differences, which seem to include unique aspects not seen in other animals. Insects are continuous (e.g., Lepidoptera and Diptera larvae) or discontinuous (e.g., predators and many hematophagous insects) feeders. The synthesis and secretion of digestive enzymes in continuous feeders seem to be constitutive, that is, they occur continuously (at least between molts), whereas in discontinuous feeders they are regulated (Lehane et al., 1996). Digestive enzymes, as with all animal proteins, are synthesized in the rough endoplasmic reticulum and processed in the Golgi

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complex, and are packed into secretory vesicles (Figure 14). There are several mechanisms by which the contents of the secretory vesicles are freed in the midgut lumen. In holocrine secretion, secretory vesicles are stored in the cytoplasm until they are released, at which time the whole secretory cell is lost to the extracellular space. During exocytic secretion, secretory vesicles fuse with the midgut cell apical membrane emptying their contents without any loss of cytoplasm (Figure 14a). In contrast, apocrine secretion involves the loss of at least 10% of the apical cytoplasm following the release of secretory vesicles (Figure 14b). These have previously undergone fusions originating larger vesicles that after release eventually free their contents by solubilization (Figure 14b). When the loss of cytoplasm is very small, the secretory mechanism is called microapocrine. Microapocrine secretion consists in releasing budding double-membrane vesicles (Figure 14c) or, at least in insect midguts, pinchedoff vesicles that may contain a single or several secretory vesicles (Figure 14d). In both cases the secretory vesicle contents are released by membrane fusion and/or by membrane solubilization caused by high pH contents or by luminal detergents.

Figure 14 Models for secretory processes of insect digestive enzymes. (a) Exocytic secretion; (b) apocrine secretion; (c) microapocrine secretion with budding vesicles; (d) microapocrine secretion with pinched-off vesicles; (e) modified exocytic secretion in hemipteran midgut cell. BSV, budding secretory vesicle; CE, cellular extrusion; DSV, double-membrane secretory vesicle; GC, Golgi complex; M, microvilli; N, nucleus; PMM, perimicrovillar membrane; PSV, pinched-off secretory vesicle; RER, rough endoplasmatic reticulum; SV, secretory vesicle. (Reprinted with permission from Terra, W.R., Ferreira, C., 2003. Digestive system. In: Resh, V.H., Carde´, R.T. (Eds.), Encyclopedia of Insects. Academic Press, San Diego, CA, pp. 313–323; ß Elsevier.)

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The secretory mechanisms of insect midgut cells reviewed below are based on immunocytolocalization data or on data combining biochemical procedures and electron micrographs. Studies based only on traditional cytology have been reviewed elsewhere (Terra and Ferreira, 1994; Lehane et al., 1996). Holocrine secretion is usually described on histological grounds mainly in midgut of insects other than higher Holometabola. These insects have large number of regenerative cells in their midguts. Thus, it is probable that cell renewal in these insects is being misinterpreted as holocrine secretion (Terra and Ferreira, 1994). In spite of this, immunocytochemical data showed that trypsin-containing vesicles along with cell organelles are discharged by opaque zone cells of adult stable flies, suggesting holocrine secretion ( Jorda˜ o et al., 1996a). Exocytic, apocrine, and microaprocrine secretory mechanisms depend largely on midgut regions. Digestive enzymes are usually secreted by exocytosis in the posterior midgut, whereas alternate mechanisms may be observed in anterior midgut. Thus, trypsin is secreted by the posterior midgut of adult mosquitoes (Graf et al., 1986), larval flies (Jorda˜ o et al., 1996b), and caterpillars (Jorda˜ o et al., 1999) by exocytosis, as well as, b-glycosidase by Tenebrio molitor middle midguts (Ferreira et al., 2002). Trypsin is secreted by the anterior midgut of caterpillars using a microapocrine route (Santos et al., 1986; Jorda˜ o et al., 1999), whereas in the anterior midgut of T. molitor amylase secretion occurs by an apocrine mechanism (Cristofoletti et al., 2001). Based only on morphological evidence, one may say that, in addition to E. ello and Spodoptera frugiperda, microapocrine secretion occurs in other lepidopteran species, such as Manduca sexta (Cioffi, 1979), whereas apocrine secretion is observed in some Orthoptera (Heinrich and Zebe, 1973) and in many coleopteran species other than T. molitor (Bayon, 1981; Silva and Souza, 1981; Baker et al., 1984). Immunocytolocalization data (Silva et al., 1995) showed that secretion by hemipteran midgut cells displays special features, as the cells have perimicrovillar membranes, in addition to microvillar ones (Figure 14e). In this case, double membrane vesicles bud from modified (double membrane) Golgi structures (Figure 14e). The double membrane vesicles move to the cell apex, their outer membranes fuse with the microvillar membrane, and their inner membranes fuse with the perimicrovillar membranes, emptying their contents (Figure 14e). Control of digestive enzyme synthesis and secretion in insects has been extensively investigated

but the results are often difficult to interpret. It is generally hypothesized that short-term variations in enzymatic activity are controlled by a secretagogue mechanism, whereas long-term changes are regulated hormonally (Lehane et al., 1996). An example of the long-term effects of hormones is the transcription of the early trypsin gene which starts a few hours after mosquito emergence and is under the control of juvenile hormone. However, the early trypsin mRNA is stored in the midgut epithelium and remains untranslated until stimulated by a secretagogue mechanism (Noriega and Wells, 1999) as described below. Other examples are known in molecular detail. A decapeptide trypsin modulating oostatic factor (TMOF) was isolated from the ovaries of A. aegypti. It is an ovarian signal that terminates trypsin biosynthesis in the midgut cells after the blood has been digested and its amino acids have been utilized for egg yolk protein synthesis (Borovsky et al., 1994). A TMOF-like factor was found in Heliothis virescens hemolymph that seems to depress trypsin synthesis at the end of each larval instar (Naven et al., 2001). The presence of food in the midgut is necessary to stimulate synthesis and secretion of digestive enzymes in some insects (Lehane et al., 1996). This secretagogue mechanism is known in molecular detail only in mosquitoes. On feeding, these insects express small amounts of early trypsin, using stored early trypsin mRNA (see above). This generates free amino acids and small peptides from blood proteins. These compounds are the initial signals that induce the synthesis and secretion of large amounts of late trypsins that complete digestion (Figure 15) (Noriega and Wells, 1999). The involvement of putative endocrine cells in the control of synthesis and secretion of digestive enzymes has frequently been proposed (Lehane et al., 1996). In support of an endocrine role for these cells, the contents of their secretory granules have been shown immunocytochemically to share epitopes with vertebrate neuropeptides. Furthermore, feeding has been shown to cause quantitative changes in the levels of these putative peptide hormones in mosquitoes (Sehnal and Zitman, 1996). However, there is no clear evidence to show what role these putative endocrine cells play in control of midgut events.

4.5.10. Concluding Remarks In spite of numerous gaps demanding further research, already indicated in this review, it is clear that insect digestive biochemistry is becoming a

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Figure 15 Model for regulation of trypsin (TRY) synthesis following a blood meal in the mosquito midgut. After emergence, juvenile hormone (JH) activates early trypsin gene transcription. Amino acids originating soon after the blood meal (unknown process) enter the cell (R1, amino acid–Naþ symporter?) and activate early trypsin translation. Early trypsin causes limited proteolysis on blood meal proteins producing unidentified peptides. These probably bind at a receptor (R2) and somehow activate late trypsin transcription. Late trypsin carries out complete proteolysis of blood meal proteins. As digestion of the blood meal nears completion, the mRNA for late trypsin disappears from the midgut. The mechanisms underlying this phenomenon are unknown (Noriega and Wells, 1999).

developed science and that its methods are powerful enough to lead to steady progress. It is conceivable that, in the next few decades, knowledge of the structural biology and function of digestive enzymes and of the control of expression of alternate digestive enzymes and their secretory mechanisms, as well as on microvillar biochemistry, will support the development of more effective and specific methods of insect control. ‘‘Quem viver, vera´ ’’–Brazilian proverb which has a similar meaning to ‘‘Whoever is alive, will see’’.

Acknowledgments The work on our laboratory greatly benefited from the long collaboration with our friend A.F. Ribeiro (Instituto de Biociencias, University of Sa˜ o Paulo) on cell biology. Also instrumental for our work were the collaboration with our friends J.R.P. Parra and M.C. Silva-Filho (Escola Superior de Agricultura Luiz de Queiroz, University of Sa˜ o Paulo) on insect nutrition and insect–plant interactions, respectively. We are indebted to Drs. J.E. Baker and B. Oppert

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(US Department of Agriculture, Agricultural Research Station, Kansas, USA), R.J. Dillon (University of Bath, UK), and S.R. Marana (University of Sa˜ o Paulo) for critical review of the manuscript. We are also grateful to our former students and now colleagues and friends A.R. Lopes and P.T. Cristofoletti for help with the literature and enzymes sequence trees and to A.B. Dias for the tree of midgut characters and for preparing all drawings. Our work was supported by Brazilian research agencies Fundac,a˜ o de Amparo a Pesquisa do Estado de Sa˜ o Paulo (FAPESP) (Tema´ tico and SMOLBnet programs) and CNPq. The authors are staff members of the Biochemistry Department and research fellows of CNPq.

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4.6 Lipid Transport D J Van der Horst, Utrecht University, Utrecht, The Netherlands R O Ryan, Children’s Hospital Oakland Research Institute, CA, USA ß 2005, Elsevier BV. All Rights Reserved.

4.6.1. Historical Perspective 4.6.1.1. Lipophorin Structure and Morphology 4.6.1.2. Lipophorin Subspecies 4.6.2. Flight-Related Processes 4.6.2.1. Adipokinetic Hormone 4.6.2.2. Strategy of the Adipokinetic Cells 4.6.2.3. Effect of Adipokinetic Hormones on Lipid Mobilization 4.6.3. Apolipophorin III 4.6.3.1. Lipid Free Helix Bundle Structure 4.6.3.2. Lipid Induced Conformation Change 4.6.3.3. Initiation of ApoLp-III Lipid Binding 4.6.3.4. ApoLp-III Alternate Functions 4.6.4. Lipophorin Receptor Interactions 4.6.4.1. The Low-Density Lipoprotein Receptor Family 4.6.4.2. Ligand Recycling Hypothesis 4.6.5. Other Lipid Binding Proteins 4.6.5.1. Lipid Transfer Particle 4.6.5.2. Carotenoid Binding Proteins 4.6.5.3. Fatty Acid Binding Proteins 4.6.5.4. Lipases 4.6.5.5. Vitellogenin

4.6.1. Historical Perspective 4.6.1.1. Lipophorin Structure and Morphology

Lipophorin was discovered 40 years ago as a major hemolymph component and key transport vehicle for water insoluble metabolites (Beenakkers et al., 1985; Chino, 1985). Lipophorin is generally regarded as a multifunctional carrier because it displays a broad ability to accommodate hydrophobic biomolecules. In essence, lipophorin can be described as a noncovalent assembly of lipids and proteins, organized as a largely spherical particle. The core of the particle is made up of hydrophobic lipid molecules, such as diacylglycerol (DAG), hydrocarbons, and carotenoids. DAG, which serves as the transport form of neutral glycerolipid in hemolymph, provides an energy source for various tissues through oxidative metabolism of its fatty acid constituents. Hydrocarbons, in the form of long-chain aliphatic alkanes and alkenes, are extremely hydrophobic lipid molecules that are deposited on the cuticle where they serve to prevent desiccation and may function as semiochemicals. Carotenoids are plant-derived pigments used for coloration and as

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a precursor to visual pigments (Canavoso et al., 2001). Another important lipid component of lipophorin is phospholipid. In general, the major glycerophospholipids present are phosphatidylcholine and phosphatidylethanolamine (Wang et al., 1992). These amphiphilic lipids exist as a monolayer at the lipophorin particle surface, positioned in such a way that their fatty acyl chains interact with the hydrophobic core of the particle while their polar head groups are presented to the aqueous milieu. In this manner, the phospholipid moieties of lipophorin serve a key structural role. The other major structural component of lipophorin is protein. All lipophorin particles possess two apolipoproteins, termed apolipophorin I (apoLp-I) and apolipophorin II (apoLp-II). ApoLp-I and apoLp-II are integral components of the lipophorin particle and cannot be removed without destruction of lipophorin particle integrity. It is recognized that apoLp-I and apoLp-II are the product of the same gene and that the two proteins arise from a posttranslational proteolytic processing event (Weers et al., 1993). This finding is consistent with the fact that apoLp-I and apoLp-II are found in a 1 : 1

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molar ratio in lipophorin particles. At present it is not known if one or the other apoLp possesses additional functions aside from its primary role in stabilizing lipophorin particle integrity although a role in receptor interactions is implied (see below). The structural role is fulfilled by the capacity of apoLp-I and apoLp-II to interact with lipid and create an interface between the nonpolar core of the particle and the external environment. In this capacity, apoLp-I and apoLp-II function in a manner similar to that proposed for apolipoprotein B in vertebrate plasma. Indeed, it is now recognized that the genes encoding these proteins are derived from a common ancestor (Babin et al., 1999). 4.6.1.2. Lipophorin Subspecies p0010

p0015

One of the hallmark features of lipophorin-mediated lipid transport relates to the dynamic nature of the particle. Lipophorin isolated from various life stages is generally of a unique density and lipid composition. For example, lipophorin from Manduca sexta fifth instar larvae displays a density of 1.15 g ml
1 with a particle diameter in the range of 16 nm. By contrast, lipophorin isolated from adult hemolymph is of lower density and larger diameter. Indeed, in M. sexta a broad array of unique lipophorin subspecies has been identified, each with characteristic properties (Prasad et al., 1986). On the basis of this diversity, a nomenclature system has been adopted that distinguishes various lipophorin subspecies based on their density. Since most particles fall within the density limits 1.21 and 1.07 g ml
1, the term high-density lipophorin (HDLp) is commonly used. Because many lipophorin subspecies are present at well-defined developmental stages, a suffix may be added to denote this. Hence, HDLp-P and HDLp-A may be used to distinguish HDLp from pupal and adult hemolymphs, respectively. One of the features of HDLp-A is its ability to associate with a third apolipophorin, apoLp-III. In insect species that use lipid as a fuel for flight (such as Locusta migratoria and M. sexta), apoLp-III is present in abundance in adult hemolymph as a lipid free protein. Whereas a small amount of apoLp-III may be associated with HDLp under resting conditions, flight activity induces association of large amounts of apoLp-III with the lipophorin particle surface (Van der Horst et al., 1979). This process, which is dependent upon the uptake of DAG by the lipophorin particle, leads to the conversion of HDLp into low-density lipophorin (LDLp). LDLp has a larger diameter, a significantly increased DAG content, and a lower density. In studies of this conversion, it has been shown that apoLp-III associates with the surface of the expanding lipophorin

particle as a function of DAG enrichment (Soulages and Wells, 1994b; Ryan and Van der Horst, 2000). Thus, it has been hypothesized that apoLp-III serves to stabilize the DAG-enriched particle, providing an interface between surface localized hydrophobic DAG molecules and the external aqueous medium. It is envisioned that continued DAG accumulation by HDLp results in partitioning of DAG between the hydrophobic core of the particle and the surface monolayer (Wang et al., 1995). The presence of DAG in the surface monolayer exerts a destabilizing effect on the particle structure and, if allowed to persist, would result in deleterious particle fusion and aggregation. By ‘‘sensing’’ the presence of DAG in the lipophorin surface monolayer, apoLp-III is attracted to the particle surface and forms a stable binding interaction. This event is fully reversible and, upon removal of DAG from the particle, apoLp-III dissociates, leading to regeneration of HDLp. Importantly, it is recognized that lipophorin particles can then bind additional DAG, forming a cycle of transport. It is noteworthy that these concepts about apoLp-III association/dissociation from lipophorin emerged from physiological studies of flight activity in L. migratoria conducted in the late 1970s and early 1980s in The Netherlands and England (Mwangi and Goldsworthy, 1977, 1981; Van der Horst et al., 1979, 1981). A cartoon depicting metabolic and biochemical processes related to induction of flight-related lipophorin conversions and the accompanying increase in neutral lipid transport capacity is presented in Figure 1. Elaboration of various aspects of this central scheme will occur in the next sections. At this point, however, it should be noted that this generalized mechanism differs fundamentally from metabolic processes in vertebrates, where lipoproteins do not have a function in the transport of energy substrates during exercise (Van der Horst et al., 2002). That being said, it is evident that novel insight into structural and functional aspects of vertebrate lipid transport processes can be gained from the study of insect lipid transport. A vivid example of this emerged recently with the identification and functional characterization of the Drosophila homolog of the vertebrate microsomal lipid transfer protein (MTP) (Sellers et al., 2003). Analysis of the Drosophila genome revealed the existence of an expressed sequence tag with 23% sequence identity to vertebrate MTP. Coexpression of this gene with that encoding a truncated version of apolipoprotein B in African green monkey kidney (COS) cells afforded the ability to assemble and secrete lipoproteins. Compared to mammalian MTP, the Drosophila MTP homolog possesses unique structural and catalytic properties that appear

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Figure 1 Molecular basis of the lipophorin lipid shuttle: AKH-controlled DAG mobilization from insect fat body during flight activity results in the reversible alternation of lipophorin from a relatively lipid-poor (HDLp) to a lipid-rich (LDLp) state, and apoLp-III from a lipid-free in a lipid-bound state. The reversible conformational change in apoLp-III induced by DAG loading of lipophorin is schematically visualized. AKHs, adipokinetic hormones; R, receptor; G, G protein; HDLp, high-density lipophorin; LDLp, low-density lipophorin; apoLp-I, -II, and -III, apolipophorin I, II, and III; TAG, triacylglycerol; DAG, diacylglycerol; FFA, free fatty acids. (Based on data from several insect species, particularly Locusta migratoria and Manduca sexta, reviewed in Ryan and Van der Horst (2000) and Van der Horst et al. (2001); and this review.)

to be related to distinct strategies of lipid transport operative in insects.

4.6.2. Flight-Related Processes 4.6.2.1. Adipokinetic Hormone

Insect flight involves the mobilization, transport, and utilization of endogenous energy reserves at extremely high rates. In insects that engage in longdistance flight, the demand for fuel, particularly lipids, by the flight muscles can remain elevated for extended periods of time. Peptide adipokinetic hormones (AKHs), synthesized and stored in neuroendocrine cells, play a crucial role in this process as they integrate flight energy metabolism. Insect AKHs are short peptides consisting of 8–11 amino acid residues. To date the structures of over 35 different AKHs are known from representatives of most insect orders; in spite of considerable variation in their structures they are clearly related (reviews: Ga¨ de, 1997; Van der Horst et al., 2001; Oudejans and Van der Horst, 2003). All AKHs are N-terminally blocked by a pyroglutamate (pGlu) residue and all but one (Ko¨ llisch et al., 2000) are C-terminally amidated. Initiation of flight activity induces the release of AKHs from the intrinsic

AKH-producing cells (adipokinetic cells) in the glandular lobes of the corpus cardiacum, a neuroendocrine gland located caudal to the insect brain and physiologically equivalent to the pituitary of mammals. The fat body plays a fundamental role in lipid storage, as well as in the process of lipolysis controlled by the AKHs. Binding of these hormones to their G protein-coupled receptors at the fat body target cells triggers a number of coordinated signal transduction processes that ultimately result in the mobilization of carbohydrate and lipid reserves as fuels for flight activity (see Figure 1). Energy-yielding metabolites are transported via the hemolymph to the contracting flight muscles. Carbohydrate (trehalose) in the circulation provides energy for the initial period of flight and is replenished from glycogen reserves. However, similar to sustained activity in many other animal species, flight activity of insects covering vast distances nonstop is powered principally by mobilization of endogenous reserves of triacylglycerol (TAG), the most concentrated form of energy available to biological tissues. As a result of TAG mobilization, the concentration of sn-1,2-DAG in the hemolymph increases progressively and gradually constitutes the principal fuel for flight. The mechanism for hormonal activation of glycogen phosphorylase, the enzyme determining

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the rate of glycogen breakdown and trehalose biosynthesis, has been well established. In contrast, little is known of the mechanism by which the pivotal enzyme TAG lipase catalyzes AKH-controlled production of the DAG on which long-distance flight is dependent. For a considerable part, the success of insects in long-distance flights is attributable to their system of neuropeptide AKHs integrating flight energy metabolism, involving the transfer of energy substrates, particularly lipids, to the flight muscles as discussed above. Therefore, recent advances in the strategy of adipokinetic cells in hormone storage and release will be discussed along with the effects of the AKHs on lipid mobilization. 4.6.2.2. Strategy of the Adipokinetic Cells

In view of their involvement in the regulation and integration of extremely intense metabolic processes, the AKH-producing cells (adipokinetic cells) of the corpus cardiacum constitute an appropriate model system for studying neuropeptide biosynthesis and processing, as well as the coherence between biosynthesis, storage, and release of these neurohormones (reviews: Ryan and Van der Horst, 2000; Van der Horst et al., 2001; Diederen et al., 2002). These processes have been particularly studied in two locust species notorious for their long-range flight capacity, L. migratoria and Schistocerca gregaria, which similarly to several other insect species mobilize more than one AKH. The three AKHs synthesized in the adipokinetic cells of L. migratoria consist of a decapeptide AKH-I and two octapeptides (AKH-II and -III). AKH-I is by far the most abundant peptide; the ratio of AKH-I : -II : -III in the corpus cardiacum is approximately 14 : 2 : 1 (Oudejans et al., 1993). All three AKHs are involved in the mobilization of both lipids and carbohydrates, although their action is differential (reviews: Vroemen et al., 1998; Van der Horst and Oudejans, 2003). In addition, several other effects of AKH are known, such as inhibition of the synthesis of proteins, fatty acids, and RNA (reviews: Ga¨de, 1996; Ga¨ de et al., 1997). The transport of these hydrophobic peptides in the circulation occurs independently of a carrier (Oudejans et al., 1996). The AKHs of L. migratoria appear to be catabolized differentially after their release; turnover half-times of AKH-I and -II during flight are relatively slow (35 and 37 min, respectively), whereas the hemolymph half-time of AKH-III is very rapid (3 min) (Oudejans et al., 1996). Degradation of the (single) AKH in the hemolymph of adult females of the cricket Gryllus bimaculatus,

which do not fly well, was estimated to be remarkably short (half-life approximately 3 min) in the resting state (Woodring et al., 2002). A recent study in which AKH concentrations were measured by radioimmunoassay shows that the hemolymph concentration of two AKHs from S. gregaria (AKH-I and -II) increases within 5 min of initiation of flight and are maintained at approximately 15-fold (AKH-I) and 6-fold (AKH-II) the resting levels over flights of at least 60 min (Candy, 2002). The increase in hormone level preceded an increase in hemolymph lipid content. Furthermore, a rapid release of the AKHs over the first few minutes was followed by a slower release, maintaining the elevated hormone levels. The AKH peptides are derived from preprohormones that are translated from separate mRNAs and subsequently enzymatically processed. Cotranslational cleavage of the signal sequences generates the AKH-I, -II, and -III prohormones, consisting of a single copy of AKH, a GKR or GRR processing site, and an AKH-associated peptide (AAP). AKH-I and -II prohormones are structurally very similar whereas AKH-III is remarkably different (Bogerd et al., 1995) (Figure 2). Prior to further processing, the AKH-I and -II prohormones dimerize at random by oxidation of their (single) cysteine residues in the AAP, giving rise to two homodimers and one heterodimer. Proteolytic processing of these dimeric products at their processing sites, involving removal of the two basic amino acid residues and amidation, using glycine as the donor, yields the bioactive hormones as well as three (two homodimeric and one heterodimeric) AKH-precursor related peptides (APRPs) with as yet unknown functions (reviews: Van der Horst et al., 2001; Diederen et al., 2002; Oudejans and Van der Horst, 2003). Recent data from capillary liquid chromatography-tandem mass spectrometry analysis indicate that these APRPs are further processed to form smaller peptides, designed AKH joining peptide 1 (AKH-JP I) and 2 (AKH-JP II), respectively (Baggerman et al., 2002) (Figure 2). The biosynthesis of AKH-III from its prohormone has only very recently been disclosed (Huybrechts et al., 2002). By the use of sophisticated techniques including capillary high-performance liquid chromatography (HPLC) and nanoflow electrospray ionization quantitative time-of-flight (Q-TOF) mass spectrometry, another (fourth) APRP was identified, a homodimer resulting from the crosslinking of two AKH-III prohormone molecules (in a parallel and/or antiparallel fashion) by two disulfide bridges formed between their (two) cysteine residues and subsequent proteolytic cleavage of the AKH-III molecules. This finding indicates that

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Figure 2 Sequence and proteolytic processing of Locusta migratoria AKH prohormones. The AKH sequence is followed by a processing site (GKR or GRR); identical residues in the AKH-associated peptides (AAPs) I and II are boxed. The cysteine residues forming disulfide bridges prior to proteolytic processing of all AKH prohormones are shown in white. JP, joining peptide. (Based on data from Bogerd, J., Kooiman, F.P., Pijnenburg, M.A.P., Hekking, L.H.P., Oudejans, R.C.H.M., et al., 1995. Molecular cloning of three distinct cDNAs, each encoding a different adipokinetic hormone precursor, of the migratory locust, Locusta migratoria: differential expression of the distinct adipokinetic hormone precursor genes during flight activity. J. Biol. Chem. 270, 23038–23043; Baggerman, G., Huybrechts, J., Clynen, E., Hens, K., Harthoorn, L., et al., 2002. New insights in adipokinetic hormone (AKH) precursor processing in Locusta migratoria obtained by capillary liquid chromatography-tandem mass spectrometry. Peptides 23, 635–644; and Huybrechts, J., Clynen, E., Baggerman, G., De Loof, A., Schoofs, L., 2002. Isolation and identification of the AKH III precursor-related peptide from Locusta migratoria. Biochem. Biophys. Res. Commun. 296, 1112–1117.)

processing of AKH-III prohormone occurs similar to that of the AKH-I and -II prohormones. In contrast to the APRPs derived from AKH-I and -II prohormones, however, no evidence was found for further processing of the APRP generated along with AKH-III production. In situ hybridization showed that the mRNA signals encoding the three different AKH preprohormones are colocalized in the cell bodies of the glandular lobes of the corpus cardiacum (Bogerd et al., 1995). Following their synthesis in the rough endoplasmic reticulum in the cell bodies, the AKH prohormones are transported to the Golgi complex and packaged into secretory granules at the transGolgi network, whereas proteolytic processing of the prohormones to bioactive AKHs is presumed to take place in the secretory granules (reviews: Van der Horst et al., 2001; Diederen et al., 2002; Oudejans and Van der Horst, 2003). The intracellular location of the AKHs was probed with antibodies specific for the corresponding associated peptides (AAP I, II, and III), the amino acid sequences of which differ to a larger degree from each other than those of the AKHs. All three (dimeric) AAPs were shown to be colocalized in the same secretory granules, which implies that these three AKHs colocalize in these granules and are released simultaneously during flight (Harthoorn et al., 1999). Since the membranes of exocytosed secretory granules fuse with the plasma membrane, the total content of the granules is released into the hemolymph. Consequently, in addition to bioactive AKHs, the APRPs, and possibly other products, are released. Whether the AKH-JPs are released is not yet clear

(Baggerman et al., 2002; Huybrechts et al., 2002); data on AKH-JP I and II indicate that these peptides do not stimulate lipid release from the fat body nor activate fat body glycogen phosphorylase, both key functions of the AKHs (Baggerman et al., 2002). The AKH cells continuously synthesize AKHs, resulting in a steady increase in the amounts of the three hormones in the corpus cardiacum with age. Concurrently, the number of the AKH-containing secretory granules (diameter 300 nm) also increases. In addition, particularly in older adults, intracisternal granules (ICGs) are produced. ICGs are present in both exocrine and endocrine cells, and originate from premature condensation of peptidergic products within cisternae of the rough endoplasmic reticulum (review: Diederen et al., 2002). In the locust adipokinetic cells, these granules, which may attain diameters up to 5 mm and even more, appear to function as a store for AKH-I and -II prohormones, as shown immunocytochemically with specific anti-AAPs (Harthoorn et al., 1999, 2000). The prohormone for AKH-III is absent, which points to differences in physiological function between AKH-III and the other two AKHs. The secretory activity of the adipokinetic cells, which has been investigated in vitro primarily for AKH-I, is subject to many regulatory substances including neurogenic locustatachykinins and humoral crustacean cardioactive peptide (CCAP) as initiating factors, trehalose as an inhibitor, and several positive and negative modulators (reviews: Van der Horst et al., 1999; Vullings et al., 1999). Recent data on the release of AKH from the corpora cardiaca in vitro show that regulatory substances

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(including CCAP) affect the release of all three AKHs in proportion to their concentration in the corpus cardiacum (Harthoorn et al., 2001). However, the only natural stimulus for the release of the AKHs is flight activity, and the relative contributions of all known substances effective in the process of release of these neurohormones remain to be established in vivo. The amount of AKHs released during flight represents only a few percent of the huge stores harbored in the adipokinetic cells. On the other hand, only a limited part of these AKH stores appear to be actually releaseable. In studies in which young secretory granules were specifically labeled, these newly formed secretory granules were preferentially released (last in, first out) (reviews: Van der Horst et al., 2001; Diederen et al., 2002; Oudejans and Van der Horst, 2003). Following the biosynthesis of new AKH prohormones, their packaging into secretory granules and their processing to bioactive AKHs, which takes less than 1 h, granules containing newly synthesized AKHs appeared to be available for release during a restricted period of approximately 8 h before they are supposed to enter a pool of older secretory granules that appear to be unable to release their content upon secretory stimulation. This indicates that only a relatively small readily releasable pool of new secretory granules exists. Therefore, an important question is whether the secretory output of AKHs during flight would induce a stimulation of the rate of AKH biosynthesis. The mRNA levels of all three AKH preprohormones, however, did not appear to be affected by flight activity, while the rate of synthesis of AKH prohormones and AKHs was not affected either (Harthoorn et al., 2001). Apparently, a coupling between release and biosynthesis of AKHs is absent. Inhibition of AKH biosynthesis in vitro by Brefeldin A, a specific blocker of the transport of newly synthesized secretory proteins from the endoplasmic reticulum to the Golgi complex, resulted in a considerable decrease in the release of AKHs induced by CCAP, and highlighted once more that the regulated secretion of AKHs is completely dependent on the existence of a readily releasable pool of newly formed secretion granules (Harthoorn et al., 2002). Therefore, we conclude that the strategy of the adipokinetic cells to cope with variations in secretory output of AKHs apparently is to rely on the continuous biosynthesis of AKHs, which produces a readily releasable pool that is sufficiently large and constantly replenished. An important question remaining unanswered is, what might be the rationale for the storage of such large quantities of hormones that are not accessible

for secretory release? In addition, the possible function of the prohormones for AKH-I and -II in the ICGs in providing an additional source of AKH prohormones when called upon remains to be established. 4.6.2.3. Effect of Adipokinetic Hormones on Lipid Mobilization

Binding of the AKHs to their plasma membrane receptor(s) at the fat body cells is the primary step to the induction of signal transduction events that ultimately lead to the activation of target key enzymes and the mobilization of lipids as a fuel for flight. Although the AKHs constitute extensively studied neurohormones and their actions have been shown to occur via G protein-coupled receptors (reviews: Van Marrewijk and Van der Horst, 1998; Vroemen et al., 1998), the general properties of which are remarkably well conserved during evolution (review: Vanden Broeck, 2001), the identification of these receptors has not been successful. Very recently, however, the first insect AKH receptors have been identified at the molecular level, namely those of the fruitfly Drosophila melanogaster and the silkworm Bombyx mori (Staubli et al., 2002). They appear to be structurally related to mammalian gonadotropin-releasing hormone (GnRH) receptors. These data promise to elucidate the nature of AKH receptors from other insects; it is envisaged that insects such as the locust, that produce two or more different types of AKH, may have two or more different AKH receptors. The signal transduction mechanism of the three locust AKHs has been studied extensively, and involves stimulation of cAMP production, which is dependent on the presence of extracellular Ca2þ. Additionally, the AKHs enhance the production of inositol phosphates including inositol 1,4,5-trisphosphate (IP3), which may mediate the mobilization of Ca2þ from intracellular stores. This depletion of Ca2þ from intracellular stores stimulates the influx of extracellular Ca2þ, indicative of the operation of a capacitative (store-operated) calcium entry mechanism. The interactions between the AKH signaling pathways ultimately result in mobilization of stored reserves as fuel for flight (reviews: Van Marrewijk and Van der Horst, 1998; Vroemen et al., 1998; Ryan and Van der Horst, 2000; Van der Horst et al., 2001; Van Marrewijk, 2003). The concentration of DAG in the hemolymph increases progressively at the expense of stored TAG reserves in the fat body, which implies hormonal activation of the key enzyme, fat body TAG lipase. In a bioassay, all three AKHs are able to stimulate lipid mobilization, although their

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relative potencies are different. This recalls the concept of a hormonally redundant system involving multiple regulatory molecules with overlapping actions (reviews: Goldsworthy et al., 1997; Vroemen et al., 1998). Results obtained with combinations of two or three AKHs, which are likely to occur together in locust hemolymph under physiological conditions in vivo, revealed that the maximal responses for the lipid-mobilizing effects were much lower than the theoretically calculated responses based on dose-response curves for the individual hormones. In the lower (probably physiological) range, however, combinations of the AKHs were more effective than the theoretical values calculated from the responses elicited by the individual hormones (review: Van Marrewijk and Van der Horst, 1998). The mechanism by which TAG lipase catalyzes AKH-controlled production of the DAG on which long-distance flight depends is only poorly understood, mainly due to technical problems in isolating or activating the lipase. In vertebrates, hormonesensitive lipase (HSL) controls mobilization of TAG stores in adipose tissue, and although contrary to insects, free fatty acids (FFA) are released into the blood for uptake and oxidation in muscle, there is a clear functional similarity between vertebrate adipose tissue HSL and insect fat body TAG lipase (reviews: Ryan and Van der Horst, 2000; Van der Horst et al., 2001; Van der Horst and Oudejans, 2003).

4.6.3. Apolipophorin III 4.6.3.1. Lipid Free Helix Bundle Structure

ApoLp-III was discovered in the late 1970s and early 1980s by research groups in Europe and North America (reviews: Blacklock and Ryan, 1994; Ryan and Van der Horst, 2000). ApoLp-III was first isolated from hemolymph of L. migratoria (Van der Horst et al., 1984) and the tobacco hawkmoth, M. sexta (Kawooya et al., 1984). Manduca sexta apoLp-III is a 166 amino acid protein that lacks tryptophan and cysteine (Cole et al., 1987). However, the well-characterized apoLp-III from L. migratoria is 164 residues long and lacks cysteine, methionine, and tyrosine (Kanost et al., 1988; Smith et al., 1994). Manduca sexta apoLp-III is nonglycosylated while L. migratoria apoLp-III contains two complex carbohydrate chains (Ha˚rd et al., 1993). Sequence analysis predicts that all apoLpIIIs are composed of predominantly amphipathic a-helix secondary structure, consistent with far ultraviolet circular dichroism (CD) studies (Ryan et al., 1993; Weers et al., 1998). An important

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breakthrough in our understanding of the structure of apoLp-III occurred with determination of the X-ray crystal structure of L. migratoria apoLp-III (Breiter et al., 1991). These authors showed that apoLp-III exists as a globular, up-and-down amphipathic a-helix bundle in the absence of lipid. The molecule is composed of five discrete a-helix segments that orient their hydrophobic faces toward the center of the bundle. Using a convenient method for bacterial overexpression, recombinant M. sexta apoLp-III was enriched with stable isotopes (Ryan et al., 1995; Wang et al., 1997b). Application of heteronuclear multidimensional nuclear magnetic resonance (NMR) techniques to isotopically enriched M. sexta apoLp-III yielded a complete assignment of this protein (Wang et al., 1997a). Structure calculations revealed a five-helix bundle molecular architecture, representing the first fulllength apolipoprotein whose high resolution solution structure has been determined in the absence of detergent (Wang et al., 2002) (Figure 3). In keeping with the X-ray structure of L. migratoria apoLp-III, this structure also reveals an up-and-down bundle of five amphipathic a-helices. Interesting, however, Wang and coworkers identified a distinct short segment of a-helix that connects helix 3 and helix 4 in the bundle (termed helix 30 ). This sequence segment (P95DVEKE100) aligns perpendicular to the long axis of the bundle and, as discussed below, has been shown to play a role in the initiation of apoLp-III lipid interaction. More recently, Fan et al. (2001, 2003) employed multidimensional NMR techniques to obtain a complete assignment and solution structure determination for L. migratoria apoLp-III. This work is significant in that it permits direct comparison between the X-ray crystal structure and the NMR structure. Interestingly, Fan et al. provide previously unreported structural evidence for a solvent exposed short helix that is positioned perpendicular to the long axis of the helix bundle. These authors propose that this short helix can serve as a recognition helix for initiation of apoLp-III lipid interaction, leading to conformational opening of the helix bundle. Contrary to the model presented by Breiter et al. (1991), Fan et al. (2003) suggest an alternate opening mechanism. Further studies will be required to elucidate the precise mechanism whereby apoLp-III recognizes and binds to available lipid surfaces (see below). 4.6.3.2. Lipid Induced Conformation Change

The up-and-down antiparallel organization of helical segments in apoLp-III allows for a simple opening of the bundle about putative ‘‘hinge’’ loops that connect the helices as originally proposed

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Figure 3 Nuclear magnetic resonance (NMR) visualizations of structure of lipid-free Manduca sexta apoLp-III. (a, b) Superposition of 40 NMR-derived structures of apoLp-III, with backbone atoms displayed in white and side chain heavy atoms displayed in green. (c) Ribbon representation of an energy-minimized, average structure of apoLp-III (PDB code 1EQ1). (Reproduced with permission from Wang, J., Sykes, B.D., Ryan, R.O., 2002. Structural basis for the conformational adaptability of apolipophorin III, a helix bundle exchangeable apolipoprotein. Proc. Natl Acad. Sci. USA 99, 1188–1193; ß by the National Academy of Sciences of the United States of America.)

by Breiter et al. (1991). The model suggests that apoLp-III initiates contact with lipid surfaces via one end of the helix bundle. Conformational opening could then occur with retention of helix boundaries present in the bundle configuration. Such an event would result in substitution of helix–helix interactions in the bundle conformation for helix– lipid interactions. Current evidence suggests that this conformational transition is triggered by availability of a suitable lipid surface and is reversible (Singh et al., 1992; Liu et al., 1993; Soulages and Wells, 1994a; Soulages et al., 1995, 1996). Thus, it is conceivable that helix 3 and helix 4 move away from helices 1, 2, and 5 in concert as the bundle opens about the loop segments connecting helix 2 and helix 3 and helix 4 and helix 5 (Breiter et al., 1991; Narayanaswami et al., 1996b). A well-known property of amphipathic exchangeable apolipoproteins in general is an ability to disrupt phospholipid bilayer vesicles and transform them into apolipoprotein–phospholipid disk complexes (Pownall et al., 1978). This property represents a useful method to investigate aspects of the proposed lipid-induced helix bundle molecular switch process. The disk-shaped complexes formed between apoLpIII and dimyristoylphosphatidylcholine (DMPC) are of uniform size and composition, permitting detailed analysis of their structural organization (Wientzek et al., 1994). Attenuated total reflectance Fourier transformed infrared spectroscopy has been employed to characterize helix orientation in apoLp-III–DMPC disk complexes (Raussens et al., 1995, 1996). This analysis, and more recent studies

(Soulages and Arrese, 2001) reveal that apoLp-III helical segments interact with phospholipid fatty acyl chains around the perimeter of the disk complex. Several independent studies have provided convincing evidence that apoLp-III undergoes a significant conformational change upon association with lipid. Kawooya et al. (1986) used a monolayer balance to investigate apoLp-III behavior at the air–water interface, while Narayanaswami et al. (1996a) studied the unique fluorescence properties of the lone tyrosine in M. sexta apoLp-III. Nearultraviolet CD analysis of L. migratoria apoLp-III indicates that helix realignment and reorientation occurs upon interaction with phospholipid vesicles (Weers et al., 1994). Sahoo et al. (2000) used pyrene excimer fluorescence spectroscopy to investigate lipid binding induced realignment of helix 2 and helix 3 in M. sexta apoLp-III. In this study cysteine residues were introduced into the protein by sitedirected mutagenesis (N40C and L90C). These sites were selected for introduction of cysteine residues based on the fact that they reside in close proximity in the helix bundle conformation. Covalent modification of the cysteine thiol groups with pyrene maleimide yielded a double pyrene labeled apoLp-III. In the absence of lipid, pyrene labeled apoLp-III adopts a helix bundle conformation. Fluorescence spectroscopy experiments revealed normal pyrene emission at 375 and 395 nm (excitation 345 nm) as well as excimer (excited state dimer) fluorescence at longer wavelengths (460 nm). Control experiments verified that the excimer peak arose from intramolecular pyrene–pyrene interactions in the

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labeled protein and was not due to intermolecular interactions. Because it is known that excimer fluorescence is manifest only when pyrene moieties are within 10 A˚ of one another, this property was used to assess the effect of lipid binding. The observation that excimer fluorescence was greatly reduced when apoLp-III was complexed with DMPC was taken as evidence for a conformational change in the protein upon lipid binding that results in relocation of helix 2 away from helix 3. In fluorescence studies of apoLp-III, carried out by Soulages and Arrese (2000a, 2000b), site-directed mutagenesis was used to create various mutant apoLp-IIIs with a single tryptophen residue in each of the five helical segments of the protein. Data obtained in this study suggests that apoLp-III undergoes a conformational change that brings helices 1, 4, and 5 into contact with the lipid surface, while others (helices 2 and 3) appear to behave differently. In other studies Soulages et al. (2001) used disulfide bond engineering to show that conformational flexibility of helices 1 and 5 of L. migratoria apoLp-III play an important role in the lipid binding process. In other studies Dettloff et al. (2001b) reported that a C-terminal truncated apoLp-III from the wax moth Galleria mellonella, comprising the first three helical segments of the protein, retains structural integrity and an ability to interact with lipid surfaces. More recently, Dettloff et al. (2002) expanded this work to encompass two additional three helix mutants derived from G. mellonella apoLp-III, a C-terminal frag-

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ment comprising helices 3–5 and a core fragment comprising helices 2– 4. All three truncation mutants retained their ability to solubilize bilayer vesicles of DMPC, an event that led to large increases in their a-helix content. The N-terminal and core fragment, but not the C-terminal fragment, were able to interact with phospholipase C modified human low-density lipoprotein, thereby preventing its aggregation. This result suggests that impairment of the lipid interaction properties of the C-terminal fragment has occurred as a result of removal of N-terminal helix segments. Taken together, it appears that the minimal essential elements required for apoLp-III lipid binding function is less than the intact five-helix bundle. Recent experiments have provided evidence that opening of the helix bundle is even more dramatic than originally postulated. It is now proposed that the protein adopts a fully extended belt-like conformation (Garda et al., 2002; Sahoo et al., 2002) (Figure 4). Garda et al. (2002) employed fluorescence resonance energy transfer methods while Sahoo et al. (2002) used pyrene excimer fluorescence to probe aspects of helix repositioning upon interaction with DMPC. In both of these studies, knowledge of the three-dimensional structure of the apoLp-III in the absence of lipid (i.e., the helix bundle conformation) allowed for structure guided site-directed mutagenesis to introduce strategically placed cysteine residues to which fluorescent reporter groups could be covalently attached. Subsequent characterization studies yielded a unifying model of apoLp-III conformation

Figure 4 Model of apoLp-III bound to phospholipid discoidal complexes. ApoLp-III complexes with phospholipids on a discoidal particle adopting an extended a-helical conformation. Lipid-triggered association involves extension of H1 away from H5, helix bundle opening and repositioning of H2 and H3. The positions of cysteine substitution mutations employed in this and previous analyses are indicated: A8C, N40C, L90C, and A138C; H1, H2, H3, and H5. Apolp-III adopts an extended helical conformation around the periphery of discoidal phospholipid bilayer complexes, with neighboring molecules aligned antiparallel with respect to each other, and shifted by one helix. (Reprinted with permission from Sahoo, D., Weers, P.M.M., Ryan, R.O., Narayanaswami, V., 2002. Lipid-triggered conformational switch of apolipophorin III helix bundle to an extended helix organization. J. Mol. Biol. 321, 201–214; ß Elsevier.)

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on disk complexes wherein the resulting structure resembles concepts and models that describe the organization of human apolipoprotein A-I on nascent high-density lipoproteins (Klon et al., 2002). 4.6.3.3. Initiation of ApoLp-III Lipid Binding

Analysis of the structure of L. migratoria apoLp-III indicates the presence of solvent exposed leucine residues at one end of the protein (Breiter et al., 1991). These authors proposed that this region of the molecule functions as a ‘‘hydrophobic sensor’’, which recognizes potential lipid surface binding sites. Surface plasmon resonance spectroscopy studies revealed that small amounts of DAG induce binding of apoLp-III to a phospholipid bilayer with its long molecular axis normal to the lipid surface (Soulages et al., 1995). This interaction is proposed to be the first step in formation of a stable binding interaction. Site-directed mutagenesis was performed to determine whether alteration in the hydrophobicity of the putative sensor region of L. migratoria apoLp-III affects its ability to initiate contact with lipid surfaces (Weers et al., 1999). In this study three partially exposed leucine residues, located at the end of the protein containing the loop segments that connect helix 1 and helix 2 and helix 3 and helix 4, were mutated to arginine. Three single arginine to leucine substitution mutants and a triple mutant were expressed in Escherichia coli and characterized in terms of their structural and stability properties. The effect of these mutations on phospholipid bilayer vesicle transformation into disk complexes versus lipoprotein binding suggests that the former binding interaction has an electrostatic component. Taken together, the data support the view that the end of the molecule bearing Leu 32, 34, and 95 is responsible for initiating contact with potential lipid surface binding sites. The solution structure of M. sexta apoLp-III revealed the presence of helix 30 at one end of the protein globule (Wang et al., 1997a, 2002). One possibility is that helix 30 reorientation facilitates contact with a lipid surface by exposing the hydrophobic interior of the helix bundle. The lipid surface could then trigger a molecular switch to induce conformational opening of the helix bundle and formation of a stable binding interaction. To investigate this, protein engineering was employed to remove helix 30 and replace it with a sequence that has a high probability of forming a b-turn (Narayanaswami et al., 1999). Characterization of the lipid binding properties of this ‘‘helix-to-turn’’ mutant apoLp-III revealed defective lipid binding properties. In more refined site-directed mutagene-

sis studies it was determined that Val 97, located in the center of helix 30 , is a critical residue for initiation of apoLp-III lipid binding. As described above, a similar short helix was identified in L. migratoria apoLp-III based on its NMR determined solution structure (Fan et al., 2003). This helix, however, is present as the opposite end of the apoLp-III helix bundle suggesting that, if it is a recognition helix, bundle opening is different from that proposed for M. sexta apoLp-III by Narayanaswami et al. (1999) and Wang et al. (2002). It is conceivable that an experimental approach similar to that employed by Narayanaswami et al. (1999) will permit direct experimental assessment of the role of this short helix in L. migratoria apoLp-III lipid interaction. Studies of the effect of the glycosyl moieties of L. migratoria apoLp-III on its lipid binding properties have also been investigated. Soulages et al. (1998) showed that recombinant apoLp-III, which lacks covalently bound carbohydrate, displayed a much stronger interaction with phospholipid vesicles than natural insect-derived apoLp-III. From the X-ray structure of L. migratoria apoLp-III in the absence of lipid, it is known that both glycosylation sites (at residues 18 and 85) are localized in the central region of the long axis of the bundle. Further study of this phenomenon revealed that apoLp-III sugar moieties interfere with helix bundle penetration into the bilayer surface during disruption and transformation into disk complexes (Weers et al., 2000). Thus, it is apparent that structural aspects of the helix bundle as well as the composition of the lipid surface influence the ability of apoLp-III to initiate and form a stable lipid-binding interaction. 4.6.3.4. ApoLp-III Alternate Functions

Based on the observed developmentally timed upregulation of its mRNA, apoLp-III has been implicated in muscle and neuron programmed cell death (Sun et al., 1995). When considered in light of its known lipid interaction properties, it is conceivable that it serves a function in membrane dissolution and/or lipid reabsorption during metamorphosis. Others have reported apoLp-III functions in insect immunity (Wiesner et al., 1997). Indeed, recent reports suggest that it is lipid associated apoLp-III that manifests this biological activity (Dettloff et al., 2001a, 2001c). These authors hypothesize that LDLp, formed in vivo, serves as an endogenous signal for immune activation, specifically mediated by lipid-associated apoLp-III interaction with hemocyte membrane receptors. From a structural standpoint, the truncated variants of G. mellonella apoLp-III (see above) that retain functional ability,

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represent useful tools to probe the structural and physiological role of apoLp-III in innate immunity. Support for this general concept has emerged from studies of G. mellonella apoLp-III variants wherein point mutations were introduced at residues 66 and 68 (Niere et al., 2001). The observation that mutation-induced decreases in apoLp-III lipid interaction properties correlate with decreased immune inducing activity is consistent with the hypothesis that apoLp-III immune activation is related to the conformational change that accompanies lipid interaction of this protein. On a broader scale it is important to understand the molecular details of this emerging group of proteins (Narayanaswami and Ryan, 2000) because the property of reversible interconversion between water-soluble and lipid bound states could have applications beyond their natural biological settings. Indeed, as work on this system continues, it is evident that apoLp-III and analogous helix bundle apolipoproteins represent novel biosurfactants with potentially useful properties, including biodegradability.

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phorin receptor (iLR; in this review LpR is adopted) shows a high similarity to mammalian LDLR. However, the ligand binding domain of LpR contains one additional cysteine-rich repeat compared to the seven repeats in LDLR, and is therefore identical to that of the human very low-density lipoprotein (VLDL) receptor (VLDLR), which also contains eight consecutive cysteine-rich repeats in this domain, as is schematically depicted in Figure 5. The amino acid sequence of the longer cytoplasmic tail of LpR is unique for insect lipophorin receptors: the 12 C-terminal amino acid residues of LDLR are completely different from those of LpR, whereas the C-terminal tail of LpR contains an additional 10 amino acid residues (Van Hoof et al., 2002). A similar VLDLR homolog was identified in mosquito oocytes and shown to bind lipophorin (Cheon et al., 2001). Three-dimensional models of the elements representing both the ligand binding

4.6.4. Lipophorin Receptor Interactions 4.6.4.1. The Low-Density Lipoprotein Receptor Family

In the concept of lipid transport during intense lipid utilization in insects, a major difference between the functioning of lipoproteins of mammals and insects is the selective mechanism by which insect lipoproteins transfer their hydrophobic cargo. Circulating HDLp particles may serve as a DAG donor or acceptor, dependent on the physiological situation, and function as a reusable lipid shuttle without additional synthesis or increased degradation of the apolipoprotein matrix, as discussed above. In apparent contrast to this concept, in fat body tissue of larval and young adult locusts, receptor-mediated uptake of HDLp was demonstrated (Dantuma et al., 1997). A receptor has been cloned and sequenced from locust fat body cDNA, and identified as a novel member of the LDL receptor (LDLR) family, that is particularly expressed in fat body, oocytes, and the brain (Dantuma et al., 1999). When stably transfected in an LDLR-deficient Chinese hamster ovary cell line, the locust receptor mediated endocytic uptake of fluorescently labeled HDLp that was absent in mock-transfected cells, suggesting that the receptor may function in vivo as an endocytic receptor for HDLp (Dantuma et al., 1999). Domain organization of this insect lipo-

Figure 5 Schematic representation of the insect lipophorin receptor (LpR) and the mammalian VLDL receptor (VLDLR), indicating the identical domain organization. The mammalian LDL receptor has the same organization, but one ligand binding repeat less. EGF, epidermal growth factor. (Based on data from Dantuma, N.P., Potters, M., De Winther, M.P.J., Tensen, C.P., Kooiman, F.P., et al., 1999. An insect homolog of the vertebrate very low density lipoprotein receptor mediates endocytosis of lipophorins. J. Lipid Res. 40, 973–978.)

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domain and the epidermal growth factor precursor homology domain of locust LpR bear a striking resemblance to those of mammalian LDLR (Van der Horst et al., 2002). Despite their pronounced structural similarity, however, the ligand specificity of LpR and LDLR for lipophorin and LDL, respectively, is mutually exclusive (Van Hoof et al., 2002). Additionally, the functioning of both receptors in lipid transport in the two animal groups seems to be intriguingly different, as discussed below. Interaction of HDLp with a specific high-affinity binding site or receptor in the cell membrane of the fat body and other tissues of several insect species has been well documented (review: Ryan and Van der Horst, 2000; see also the recent data by Pontes et al., 2002; Van Hoof et al., 2003). In the binding of human LDL to its receptor, the most C-terminal 1000 amino acids in apoB are involved (Bore´ n et al., 1998). Remarkably, although both the sequence and domain structure of the precursor protein of insect apoLp-I and apoLp-II (apoLp-II/I) resemble that of apoB100 (Babin et al., 1999; Van der Horst et al., 2002), apoLp-II/I does not show homology to this C-terminal part of apoB100, leaving the receptor-binding domain of apoLp-II/I to be disclosed in the future. Immunocytochemical localization of HDLp has demonstrated the presence of the lipoprotein in endosomes of fat body of the larval dragonfly Ashna cyanea (Bauerfeind and Komnick, 1992) and in developing mosquito oocytes (Sun et al., 2000), suggesting endocytosis of circulating HDLp. Uptake of HDLp in the fat body of young adult locusts was shown to be receptor mediated (Dantuma et al., 1997). A recent study presents evidence for the involvement of LpR in the endocytic uptake mechanism for HDLp in the locust that is temporally present during specific periods of development (Van Hoof et al., 2003). Shortly after ecdysis, when lipid reserves are depleted, LpR is expressed in fat body tissue of young adult locusts as well as larvae, and fat body cells are able to endocytose the complete HDLp particle. On the fourth day after (larval or imaginal) ecdysis, however, expression of LpR is downregulated and drops below detectable levels; consequently, HDLp is no longer internalized. Downregulation of LpR was postponed by experimental starvation of adult locusts immediately after ecdysis. Moreover, by starving adult locusts after downregulation of LpR, expression of the receptor was induced. These data suggest that LpR expression is regulated by the demand of fat body tissue for lipid components (Van Hoof et al., 2003).

4.6.4.2. Ligand Recycling Hypothesis

Receptor-mediated uptake of HDLp in newly ecdysed adult and larval locusts may provide a mechanism for the uptake of specific lipid components separate from the mechanism of selective unloading of HDLp lipid cargo at the cell surface. At this time, however, the simultaneous existence of the two distinct mechanisms cannot be excluded. Downregulation of LpR in fat body cells suggests that this receptor is not involved in the lipophorin shuttle mechanism operative in the flying insect. Nevertheless, an endocytic uptake of HDLp seems to conflict with the selective process of lipid transport between HDLp and fat body cells without degradation of the lipophorin matrix. However, the pathway followed by the internalized HDLp may be different from the classical receptor-mediated lysosomal pathway typical of LDLR-internalized ligands. Therefore, an intriguing question is whether this novel LpR, in contrast to all other members of the LDLR family, is able to recycle its ligand after intracellular trafficking. In mammalian cells, LDL and diferric transferrin have been used extensively to study intracellular transport of ligands that are internalized by receptor-mediated endocytosis (Goldstein et al., 1985; Brown and Goldstein, 1986; Mellman, 1996; Mukherjee et al., 1997). Whereas LDL dissociates from its receptor and is completely degraded in lysosomes, transferrin remains attached to its receptor and is eventually resecreted from the cells (Ghosh et al., 1994). LDLRexpressing Chinese hamster ovary cells transfected with LpR cDNA were used to study the endocytic uptake and intracellular pathways of locust HDLp simultaneously with either human LDL or transferrin. Intracellular trafficking of multiple fluorescently labeled ligands was visualized by multicolor confocal laser scanning microscopy, and provided evidence for different intracellular routes followed by the mammalian and insect lipoproteins (Van Hoof et al., 2002) (Figure 6). In contrast to LDL, which is degraded in lysosomes after dissociating from its receptor, HDLp remained coupled to LpR and was transported to a nonlysosomal juxtanuclear compartment. Colocalization of HDLp with transferrin identified this organelle as the endocytic recycling compartment, from which internalized HDLp was eventually resecreted (half-time 13 min), in a manner similar to that operative in the transferrin recycling pathway. The above data indicate that, in mammalian cells, endocytosed insect HDLp, in contrast to human LDL, follows a recycling pathway mediated by LpR. This promises to elucidate new aspects of

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Figure 6 Confocal laser microscopic digital image of Chinese hamster ovary cells incubated with fluorescently labeled HDLp (a) and transferrin (b) after a chase period of 20 min. Colocalization of both ligands is visualized in yellow when images (a) and (b) are merged (c). (Reproduced with permission from Van der Horst, D.J., Van Hoof, D., Van Marrewijk, W.J.A., Rodenburg, K.W., 2002. Alternative lipid mobilization: the insect shuttle system. Mol. Cell. Biochem. 239, 113–119; ß Kluwer Academic Publishers.)

LDLR functions, although recycling of endocytosed HDLp in insect fat body cells remains to be shown. Additionally, although the acidic endosomal environment of endocytosed HDLp has been postulated to facilitate the transfer of lipid components other than DAG or cholesterol (Dantuma et al., 1997), both the function of the process of receptormediated endocytosis and the rationale for its occurrence during specific stages of insect development remain to be elucidated.

4.6.5. Other Lipid Binding Proteins 4.6.5.1. Lipid Transfer Particle

The concept that specialized proteins exist, which function in redistribution of hydrophobic lipid molecules, is well documented in the mammalian literature. A wide variety of distinct lipid transfer proteins have been characterized and their metabolic roles investigated. In 1986 a lipid transfer particle (LTP) was isolated from M. sexta larvae and shown to facilitate vectorial redistribution of lipids among plasma lipophorin subspecies (Ryan et al., 1986a, 1986b). In subsequent studies LTP was implicated in formation of LDLp from HDLp in response to AKH (Van Heusden and Law, 1989). The concept that LTP functions in flight-related lipophorin conversions correlates well with observed increases in LTP concentration in adult hemolymph compared with other developmental stages (Van Heusden et al., 1996; Tsuchida et al., 1998). When compared to other lipid transfer proteins, however, LTP displays unique structural characteristics. For example, it exists as a high molecular weight complex of three apoproteins (apoLTP-I, 320 000 kDa; apoLTP-II, 85 000 kDa; and apoLTP-III, 55 000 kDa) and 14% noncovalently associated lipid (Ryan et al., 1988). LTPs exhibiting similar structural properties have

been isolated from L. migratoria, Periplaneta americana, B. mori, and Rhodnius prolixus hemolymph (Hirayama and Chino, 1990; Takeuchi and Chino, 1993; Tsuchida et al., 1997; Golodne et al., 2001). The large size of LTP permitted examination of its structural properties by negative stain electron microscopy (Ryan et al., 1990a; Takeuchi and Chino, 1993). LTP from these two distinct species displays a highly asymmetric morphology with two major structural features, a quasispherical head region and an elongated cylindrical tail, which appears to possess a central hinge. The lipid component resembles that of lipophorin in that it contains predominantly phospholipid and DAG (Ryan, 1990). An important question arising from these physical characteristics relates to the requirement of the lipid component as a structural entity and/or its involvement in catalysis of lipid transfer. Studies employing lipoproteins containing radiolabeled lipids in incubations with LTP have revealed that the lipid component of the particle is in dynamic equilibrium with that of lipoprotein substrates (Ryan et al., 1988). Thus, it is evident that the lipid moiety is not merely a static structural component of LTP. Rather, it can be considered as a functional element in the mechanism of lipid transfer. Very recently, studies of the Drosophila genome revealed the presence of a homolog of mammalian microsomal lipid transfer protein, MTP. Sellers et al. (2003) provided evidence that Drosophila MTP possesses the functional ability to transfer TAG onto nascent apolipoprotein B-containing lipoproteins in transfected COS cells, although it does not facilitate transfer of TAG among vesicles in vitro. Future studies, designed to elucidate the physiological role of this newly identified insect MTP and its participation in lipophorin assembly,

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interconversions or some other aspect of lipid transport, represents a fruitful direction for future research. Given the size similarity between Drosophila MTP and apoLTP-II from other insect species, as well as their apparent functional similarities (see below), it is conceivable that apoLTP-II may be related to insect MTP. 4.6.5.1.1. Lipid substrate specificity Experiments have been conducted to examine the ability of LTP to utilize various substrate lipids. As reviewed earlier (Ryan and Van der Horst, 2000; Arrese et al., 2001) LTP catalyses the exchange and net transfer of DAG, in keeping with its proposed role in lipophorin interconversions in vivo. Extending this concept, Canavoso and Wells (2001) incubated radiolabeled midgut sacs with lipophorin containing medium. These authors found that transfer of DAG from the midgut sacs to lipophorin was blocked by preincubation with antibody against LTP, supporting the view that LTP functions in DAG export from the midgut to lipophorin. In a similar manner, LTP was shown by Jouni et al. (2003) to be required for DAG transfer from lipophorin to B. mori ovarioles. In studies of other potential lipid substrates, Singh and Ryan (1991) used [14C1]acetate to label the DAG and hydrocarbon moiety of lipophorin in vivo. Subsequent lipid transfer experiments revealed that LTP is capable of facilitating transfer of hydrocarbon among lipoprotein substrates, suggesting LTP plays a role in movement of these extremely hydrophobic, specialized lipids from their site of synthesis to their site of deposition at the cuticle (Takeuchi and Chino, 1993). Interestingly, the rate of LTP-mediated hydrocarbon transfer was slower than DAG transfer. In other work, B. mori LTP was employed in studies of LTP-mediated carotene transfer among lipophorin particles (Tsuchida et al., 1998). Again, compared to DAG transfer, the rate of LTP-mediated carotene redistribution was much slower. Taken together, these results suggest that LTP may have a preference for DAG versus hydrocarbon or carotenes. Alternatively, the observed preference for DAG may be a function of the relative accessibility of the substrates within the donor lipoprotein. The ability of LTP to facilitate phospholipid transfer was studied by Golodne et al. (2001). These authors observed that LTP mediated a time-dependent transfer of phospholipid that was nonselective. In contrast to the requirement for LTP to mediate transfer of DAG, hydrocarbon, carotenoids, and phospholipids, Yun et al. (2002) provided evidence that LTP does not function in cholesterol transfer or redistribution in

M. sexta. Rather, cholesterol is proposed to diffuse among tissues via mass action, freely transferring between lipophorin and tissues, depending on the physiological need. In keeping with this interpretation, Jouni et al. (2003) found that cholesterol transfer from lipophorin to B. mori ovarioles was unaffected by antibodies directed against LTP, whereas DAG transfer was inhibited. 4.6.5.1.2. Mechanism of facilitated lipid transfer In general lipid transfer catalysts may act as carriers of lipid between donor and acceptor lipoproteins or transfer may require formation of a ternary complex between donor, acceptor, and LTP. Based on the observed LTP-mediated net transfer of DAG from HDLp to human LDL (Ryan et al., 1990b), a strategy was developed to address this question experimentally (Blacklock et al., 1992). [3H]-DAG–HDLp and unlabeled LDL were covalently bound to Sepharose matrices and packed into separate columns connected in series, followed by circulation of LTP or buffer. Circulation of LTP, but not buffer, resulted in a concentrationdependent increase in the amount of radiolabeled DAG recovered in the LDL fraction, revealing that LTP facilitates net lipid transfer via a carriermediated mechanism. Blacklock and Ryan (1995) employed LTP apolipoprotein specific antibodies to probe the structure and catalytic properties of M. sexta LTP, obtaining evidence that apoLTP-II is a catalytically important apoprotein. In a similar manner Van Heusden et al. (1996) employed LTP antibody inhibition experiments to demonstrate that all three LTP apoproteins are important for lipid transfer activity. These authors found that, unlike apoLp-III, apoLTP-III is not found as a free protein in hemolymph (Van Heusden et al., 1996) despite the fact that it dissociates from the complex following exposure to nonionic detergent (Blacklock and Ryan, 1995). 4.6.5.2. Carotenoid Binding Proteins

In insects, the involvement of lipophorin in the hemolymph transport of dietary carotenoids is well documented (Tsuchida et al., 1998). Lipophorin may selectively deposit these isoprenoid lipid components at specific tissues, which is in keeping with the general function of the insect lipoprotein to selectively accept and deliver its hydrophobic cargo. The mechanism involved in this selective delivery is not fully understood, and may involve LTP and/or the lipophorin receptor, as discussed above. Carotenoids fulfill several important roles in insects. Certain carotenoids are the provitamins for vitamin A, which is required as the visual

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pigment chromophore in the animal kingdom, including D. melanogaster (Giovannucci and Stephenson, 1999). Another example is the production of a yellow cocoon in the silkworm, B. mori, which is dependent on the availability of carotenoids in the silk gland (Tabunoki et al., 2002). On both aspects, novel data illustrate both the presence and function of carotenoid binding proteins in insects. Recently, a carotenoid binding protein (CBP) from the silk gland of B. mori larvae has been identified (Tabunoki et al., 2002). The function of the 33 kDa protein was studied using several phenotypes of B. mori mutants; only in mutants carrying the dominant Y (yellow hemolymph) gene CBP is present in the villi of the midgut epithelium, suggesting that CBP may be involved in absorption of carotenoids. Similarly, using another mutant, it was inferred that CBP is involved in the uptake of carotenoids by the silk gland. The deduced amino acid sequence from CBP cDNA revealed the protein to be a novel member of the steroidogenic acute regulatory (StAR) protein family; CBP binds carotenoids rather than cholesterol. Another protein involved in the cellular uptake of dietary carotenoids was discovered in Drosophila. Recent studies demonstrate that the molecular basis for blindness of the ninaD visual mutant fly is a defect in the cellular uptake of carotenoids (Kiefer et al., 2002). The ninaD gene encodes a class B scavenger receptor with significant sequence identity to the mammalian class B scavenger receptors, SR-BI and CD36; the loss of this function abolishes carotenoid uptake and results in a vitamin A-deficient phenotype. In mammals, class B scavenger receptors, particularly SR-BI, are involved in cholesterol homeostasis and mediate the bidirectional flux of unesterified cholesterol between target cells and lipoproteins ( Jian et al., 1998; Yancey et al., 2000). As lipophorin is structurally related to mammalian lipoproteins, it was speculated that the ninaD scavenger receptor mediates the transfer of carotenoid from lipophorin in a mechanistically similar manner (Kiefer et al., 2002). ninaD mRNA levels were particularly high in pupae, which may indicate that NinaD has an important role in the redistribution of zeaxanthin, the larval storage form of carotenoids in the fat body, to the developing eyes during pupation (Giovannucci and Stephenson, 1999). Interestingly, in 1–2-day-old adults, ninaD mRNA levels were equally high, in contrast to older adults (>10 days) where ninaD mRNA was hardly visible. This suggests a similarity to the expression of the lipophorin receptor in the adult locust (Van Hoof et al., 2003), and may indicate that both proteins function in

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specific lipid metabolic processes in the young adult stage that are no longer required in the older adult. 4.6.5.3. Fatty Acid Binding Proteins

Hydrolysis of LDLp-carried DAG by a lipophorin lipase at the flight muscles (review: Van der Horst et al., 2001) results in the extracellular production of FFAs. After uptake by the flight muscle cells, these FFAs are oxidized for energy generation. The mechanism by which the extracellularly liberated FFAs are translocated across the plasma membrane is as yet unknown, but may involve membrane fatty acid transporter proteins similar to those identified in mammals, i.e., fatty acid translocase (FAT)/CD36, fatty acid transport protein (FATP), and plasma membrane fatty acid binding protein (FABPpm) (reviews: Glatz and Van der Vusse, 1996; Brinkmann et al., 2002). The intracellular transport of FFAs in insect flight muscle is mediated by a fatty acid binding protein (FABP) (review: Haunerland, 1997). This insect FABP belongs to the cytoplasmic FABPs that comprise a family of 14–15 kDa proteins that bind fatty acid ligands with high affinity and are involved in shuttling fatty acids to cellular compartments, modulating intracellular lipid metabolism, and regulating gene expression (review: Boord et al., 2002). Intriguingly, in contrast to FABP in mammals, locust FABP is an adult-specific protein, the expression of which is directly linked to metamorphosis; to accomplish the extremely high metabolic rate encountered during migratory flight, the concentration of FABP in locust flight muscle cytosol is over three times that as in the mammalian heart (review: Haunerland, 1997). The high amino acid sequence similarity (82%) between the FABP of L. migratoria flight muscle and that of human skeletal muscles (Maatman et al., 1994) is reflected in a strong similarity in their three-dimensional structures (Van der Horst et al., 2002). 4.6.5.4. Lipases

Similar to those in mammals, the lipids that are oxidized during sustained physical exercise in insects are derived from stored TAG reserves by the action of a lipase. In mammals, however, FFAs are mobilized from adipose tissue and transported in the blood bound to serum albumin for uptake and oxidation in muscle, whereas in insects, DAG is released from the fat body and transported to the flight muscles by lipophorin as discussed above, requiring hydrolysis of DAG at the flight muscles to liberate the fatty acids that are eventually taken up. The lipase catalyzing the hydrolysis of TAG

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stores in both mammals and insects plays a crucial role in energy metabolism by providing the source of energy for the working muscles, and thus between mammalian adipose tissue lipase (HSL) and insect fat body TAG lipase there is a clear functional similarity. However, there is an essential difference in the mode of action of both lipolytic enzymes, as TAG hydrolysis by HSL results in the release of FFAs, whereas that of the insect TAG lipase results in release of DAG. HSL is regulated posttranscriptionally by phosphorylation; structural studies of the enzyme have identified several amino acids and regions of the molecule that are critical for enzymatic activity and regulation of HSL (reviews: Holm et al., 1997; Kraemer and Shen, 2002). In addition, other lipiddroplet associated proteins such as perilipins and lipoptransin may be involved in the interaction of HSL with the stored lipid (reviews: Ryan and Van der Horst, 2000; Van der Horst et al., 2001). Recent data in which the acute changes in HSL activity of human adipose tissue were monitored during physical exercise by assaying the phosphorylation state of the enzyme show that lipase activity increased over sixfold above baseline at 5 min of exercise and subsequently decreased gradually despite continuation of the exercise stimulus (Petridou and Mougios, 2002). 4.6.5.4.1. Regulation of lipolysis In spite of the importance of lipid mobilization for sustained insect flight, the regulation of lipolysis in insect fat body is largely unknown. The involvement of AKHs in lipolysis is without debate and was demonstrated both in vivo from enhanced levels of DAG in hemolymph of insects injected with the hormones and in vitro by the accumulation of DAG in isolated L. migratoria fat body tissue that was incubated in the presence of AKH; both cAMP and Ca2þ were shown to play an important role in the effect of AKH on lipolysis (reviews: Ryan and Van der Horst, 2000; Van der Horst et al., 2001; Van der Horst and Oudejans, 2003). In two insect species that rely on lipid mobilization during flight activity, L. migratoria and M. sexta, it has been shown that the DAG, which is released from the fat body by the action of AKH, is stereospecific and has the sn-1,2 configuration (review: Ryan and Van der Horst, 2000). The pathway for the stereospecific synthesis of this sn-1,2-DAG is not well understood. Although a number of experimental data suggest that a stereospecific TAG lipase is involved (Arrese and Wells, 1997; Arrese et al., 2001) other data obtained by the same authors are conflicting with this view

(review: Ryan and Van der Horst, 2000). For instance, direct stereospecific hydrolysis of TAG into sn-1,2-DAG would involve enzymatic removal of the sn-3 fatty acid. As the latter fatty acid was not accumulated in the fat body nor in the hemolymph (Arrese et al., 2001), conclusive data demonstrating unambiguously the occurrence of this mode of TAG lipase action is still lacking. In summary, unlike the functional similarity between HSL in vertebrate adipose tissue and TAG lipase in insect fat body, there is a discrepancy in the mode of action of both lipolytic enzymes. The vertebrate HSL catalyzes nonstereospecific hydrolysis of TAG (as well as of DAG and monoacylglycerol) (review: Kraemer and Shen, 2002), leading to the release of fatty acids, whereas the action of the insect TAG lipase results in the formation and release of sn-1,2-DAG. So far, the reason for this discrepancy is unknown, but both structural and regulatory aspects may be involved. Elucidation of the mechanism of TAG lipase action and the regulation of its activity is essential for a better insight in the as yet poorly understood mechanism of flight-directed lipolysis in insects. 4.6.5.5. Vitellogenin

Oocyte development in adult females involves the accumulation of large amounts of lipid, most of which is extraovarian in origin and is delivered by lipophorin. Another lipid binding protein that serves a role in lipid transport to the oocyte is vitellogenin; although its overall contribution to the oocyte lipid accumulation is relatively modest (about 5%) (Sun et al., 2000). While the structural properties of insect vitellogenins are diverse, they generally possess 10% lipid, primarily phospholipid and glycerolipid. Vitellogenin is synthesized and assembled in the fat body, secreted into hemolymph, and taken up by oocytes. Vitellogenin uptake is facilitated by members of a subfamily of the LDLR family that have been characterized in Drosophila (Schonbaum et al., 1995) and in the mosquito Aedes aegypti (Sappington et al., 1996). Recently, it was demonstrated that the ovarian vitellogenin receptor (VgR) is only distantly related to the lipophorin receptor (LpR), another ovarian LDLR homolog with a different ligand (Cheon et al. 2001). These data imply that the receptormediated mechanisms involved in the uptake of lipid and the accumulation of yolk protein precursors (which provide a key nutrient source for the developing oocyte), utilize two separate receptors (VgR and LpR) which are specific for their respective ligands, vitellogenin and lipophorin. Considerable early work was performed to characterize

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the lipid transport properties of vitellogenin (review: Kunkel and Nordin, 1985), while more recent work has focused on molecular and evolutionary aspects of vitellogenin proteins and receptor-mediated endocytosis. These aspects, which are beyond the scope of the present treatise, have been comprehensively reviewed elsewhere (Sappington and Raikhel, 1998; Raikhel et al., 2002) With respect to lipophorin internalization, it is important to note that immunocytochemical data in the mosquito (Sun et al., 2000) revealed that only a small amount of lipophorin accumulates in developing oocytes as yolk protein, comprising 3% of total ovarian proteins upon completion of protein internalization. Since lipid accounts for 35–40% of the insect egg dry weight (Kawooya and Law, 1988), Sun et al. (2000) proposed that internalization of lipophorin is unlikely to be the major route of lipid delivery to the developing oocyte. A dual mechanism for lipophorin-mediated lipid delivery to oocytes (a lipophorin shuttle mechanism involving LDLp and internalization of HDLp, with stripping of most of its lipid) has been demonstrated earlier (Kawooya and Law, 1988; Kawooya et al., 1988; Liu and Ryan, 1991). However, considering that the LpR receptor involved in uptake of HDLp by mosquito oocytes bears a high structural similarity to the LpR discovered in locust fat body cell membranes (Dantuma et al., 1999; Cheon et al., 2001), the precise mechanism of LpR-mediated endocytosis in the oocyte, and the fate of HDLp, remain open questions. In light of the ligand recycling hypothesis discussed earlier for lipid delivery to the fat body, the possibility exists that LpR recycles its ligand after intracellular trafficking, providing another mechanism for the uptake of specific lipid components by the oocyte.

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Weers, P.M.M., Van der Horst, D.J., Ryan, R.O., 2000. Interaction of locust apolipophorin III with lipoproteins and phospholipid vesicles: effect of glycosylation. J. Lipid Res. 41, 416–423. Weers, P.M.M., Van Marrewijk, W.J.A., Beenakkers, A.M.T., Van der Horst, D.J., 1993. Biosynthesis of locust lipophorin: apolipophorins I and II originate from a common precursor. J. Biol. Chem. 268, 4300–4303. Weers, P.M.M., Wang, J., Van der Horst, D.J., Kay, C.M., Sykes, B.D., et al., 1998. Recombinant locust apolipophorin III: characterization and NMR spectroscopy. Biochim. Biophys. Acta 1393, 99–107. Wientzek, M., Kay, C.M., Oikawa, K., Ryan, R.O., 1994. Binding of insect apolipophorin III to dimyristoylphosphatidylcholine vesicles: evidence for a confirmational change. J. Biol. Chem. 269, 4605–4612. Wiesner, A., Losen, S., Kopacek, P., Weise, C., Gotz, P., 1997. Isolated apolipophorin III from Galleria mellonella stimulates the immune reactions of this insect. J. Insect Physiol. 43, 383–391. Woodring, J., Lorenz, M.W., Hoffmann, K.H., 2002. Sensitivity of larval and adult crickets (Gryllus bimaculatus) to adipokinetic hormone. Comp. Biochem. Physiol. A 133, 637–644. Yancey, P.G., de la Llera-Moya, M., Swarnakar, S., Monzo, P., Klein, S.M., et al., 2000. High density lipoprotein phospholipid composition is a major determinant of the bi-directional flux and net movement of cellular free cholesterol mediated by scavenger receptor BI. J. Biol. Chem. 275, 36596–36604. Yun, H.K., Jouni, Z.E., Wells, M.A., 2002. Characterization of cholesterol transport from midgut to fat body in Manduca sexta larvae. Insect Biochem. Mol. Biol. 32, 1151–1158.

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4.7 Proteases M R Kanost and T E Clarke, Kansas State University, Manhattan, KS, USA ß 2005, Elsevier BV. All Rights Reserved.

4.7.1. Introduction and History 4.7.2. Proteases in Eggs and Embryos 4.7.2.1. Proteases That Digest Egg Yolk Proteins 4.7.2.2. The Dorsal Pathway in Embryonic Development 4.7.3. Hemolymph Plasma Proteases 4.7.3.1. Serine Proteases 4.7.3.2. Protease Inhibitors 4.7.4. Cellular Proteases 4.7.4.1. Serine Proteases 4.7.4.2. Cathepsin-Type Cysteine Proteases 4.7.4.3. Caspases 4.7.4.4. Metalloproteases 4.7.4.5. Aspartic Acid Proteases 4.7.4.6. Proteasomes 4.7.5. Conclusions and Future Prospects

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4.7.1. Introduction and History

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Proteases (peptidases) are enzymes that hydrolyze peptide bonds in proteins. Exopeptidases cleave a terminal amino acid residue at the end of a polypeptide; endopeptidases cleave internal peptide bonds. Hooper (2002) provides a useful introduction to the general properties of proteases. Proteases can be classified based on the chemical groups that function in catalysis. In serine proteases the hydroxyl group in the side chain of a serine residue in the active site acts as a nucleophile in the reaction that hydrolyzes a peptide bond, whereas in cysteine proteases the sulfhydryl group of a cysteine side chain performs this function. In aspartic acid proteases and metalloproteases, a water molecule in the active site (positioned by interacting with an aspartyl group or a metal ion, respectively) functions as the nucleophile that attacks the peptide bond. Proteases are classified on this basis of catalytic mechanism in a system developed by the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (http://www.chem.qmul.ac.uk/ iubmb/enzyme/EC3/4/). However, proteases can have the same catalytic mechanism but will be unrelated in amino acid sequence, as products of convergent evolution. The MEROPS classification system groups proteases into families based on sequence similarity (Rawlings and Barrett, 1999) (http://merops.sanger.ac.uk). A protease cleaves a peptide bond, called the scissile bond, between two amino acid residues named

P1 and P10 (Schechter and Berger, 1967). Residues on the N-terminal side of the scissile bond are numbered in the C to N direction, whereas residues on the C-terminal side of the scissile bond (the ‘‘prime’’ side) are numbered in the N to C direction (Figure 1). The substrate specificity of most endopeptidases is highly dependent on the nature of the side chain of the P1 residue, but the sequence of other residues near the scissile bond can also affect binding of the substrate to the active site and thus influence substrate specificity. Insects produce abundant proteases that function in digestion of dietary proteins in the gut. Such proteases are thoroughly discussed in 00053. This chapter focuses on nondigestive proteases, which have many diverse roles in insect biology. These proteases often function in cascade pathways, in which one protease activates the zymogen form of another protease, leading to amplification of an initial signal that may involve a few molecules and finally generating a very large number of effector molecules at the end of the pathway. The complement and blood coagulation pathways in mammals are well-understood examples of this type of protease cascade, which also occur in insect embryonic development and insect immune responses (Jiang and Kanost, 2000; Krem and DiCerra, 2002; see Chapter 4.5). Details of the organization and regulation of such pathways in insect biology are beginning to be understood. This chapter will include an

248 Proteases

Figure 1 The Schechter and Berger (1967) notation for protease cleavage sites. The arrow designates the scissile peptide bond between amino acid residues P1 and P10 .

emphasis on the current state of knowledge in this rapidly developing area. Insect proteases have previously been reviewed by Law et al. (1977), Applebaum (1985), Terra et al. (1996), and Reeck et al. (1999). These reviews deal primarily with proteases as they function in the digestion of food. Only quite recently has much detailed information appeared about proteases with other functions in insect biology. An exception is cocoonase, the first insect protease that was purified and well-characterized biochemically. Cocoonase is a serine protease from silk moths that functions to hydrolyze silk proteins in the cocoon, enabling the adult moth to emerge (Kafatos et al., 1967a, 1967b). It digests sericin, the silk protein that cements fibroin threads together (see Chapter 2.11). A specialized tissue called the galea, derived from modified mouthparts, synthesizes and secretes the zymogen form, prococoonase (Kafatos, 1972). On the surface of the galea, prococoonase is activated by cleavage at a specific site by an unknown protease in the molting fluid (Berger et al., 1971). Sequencing of an amino terminal fragment and the peptide containing the active site Ser residue indicated that the activation and catalytic mechanisms of coccoonase were quite similar to those of mammalian trypsin (Felsted et al., 1973; Kramer et al., 1973). It is surprising that no molecular cloning has apparently yet been carried out for this historically important insect protease.

4.7.2. Proteases in Eggs and Embryos 4.7.2.1. Proteases That Digest Egg Yolk Proteins

Vitellin and a few other egg-specific proteins stored in yolk granules of insect eggs are digested by proteases to release amino acids for use in embryonic development (Raikhel and Dhadialla, 1992; Chapter 3.9). Such enzymes in eggs represent several different protease families and mechanistic classes. In Bombyx mori, a serine protease that degrades vitellin was purified from B. mori eggs (Indrasith et al., 1988), and its cDNA was cloned (Ikeda et al., 1991). This protease cleaves after Arg or Lys P1 residues and is a member of the S1

(chymotrypsin-like) family of serine proteases. It is synthesized in ovaries as a zymogen and is activated during embryogenesis. A second serine protease from the S1 family specifically degrades the 30 kDa yolk proteins present in B. mori eggs (Maki and Yamashita, 1997, 2001). This protease, which is synthesized at the end of embryogenesis, has elastase-like specificity, cleaving after P1 residues with small side chains. A serine carboxypeptidase is synthesized in the fat body of a mosquito, Aedes aegypti, transported through the hemolymph, and taken up by oocytes (Cho et al., 1991). This protease is synthesized as a zymogen and activated within eggs during embryogenesis. Cysteine proteases have been characterized from eggs of several insect species. Those that have been sequenced are from the C1 (papain-like) family of cysteine proteases. They typically have acidic pH optima and have biochemical properties similar to mammalian cysteine proteases known as cathepsins (although not all proteases called cathepsins are cysteine proteases). A 47 kDa cysteine protease that can digest vitelllin has been purified from B. mori eggs (Kageyama and Takahashi, 1990; Yamamoto and Takahashi, 1993), and its cDNA has been cloned (Yamamoto et al., 1994). It has sequence similarity to mammalian cathepsin L and a preference for cleaving at sites that have hydrophobic residues at the P2 and P3 positions. It is synthesized as a zymogen in ovary and fat body as a maternal product and taken up into oocytes (Yamamoto et al., 2000). This cysteine protease is self-activated at low pH by proteolytic processing, apparently by a weak activity of the proenzyme under acidic conditions (Takahashi et al., 1993). Cysteine proteases with sequence similarity to mammalian cathepsin B have also been identified as enzymes that digest insect egg yolk proteins. In Drosophila melanogaster, a cysteine protease is associated with yolk granules (Medina et al., 1988). Its zymogen is apparently activated by a serine protease during embryonic development, and the active cathepsin-B then digests yolk proteins. A cysteine protease that digests yolk proteins has also been identified in another higher dipteran, Musca domestica (Ribolla and De Bianchi, 1995). In A. aegypti a ‘‘vitellogenic cathepsin B’’ is synthesized in adult female fat body after the insect has taken a blood meal, and the zymogen is transported through the hemolymph and taken up by oocytes (Cho et al., 1999). The enzyme is activated by proteolytic processing when embryonic development begins and then probably functions to digest vitellin. A cathepsin B-like protease that can digest vitellin is also synthesized in the fat body and ovaries of a

Proteases

lepidopteran insect, Helicoverpa armigera (Zhao et al., 2002). However, its gene is expressed in fat body of males and females and in larvae and pupae, and thus is not coordinated with vitellogenesis as is the mosquito cathepsin-B. 4.7.2.2. The Dorsal Pathway in Embryonic Development

A signal transduction system that regulates dorsal/ ventral patterning in D. melanogaster embryonic development is activated by an extracellular serine protease cascade (Morisato and Anderson, 1995; LeMosy et al., 1999). The members of this cascade are produced maternally and deposited in the space between the vitellin membrane and the embryo. The pathway was elucidated by genetic analysis, and recently the recombinant forms of the proteases have been studied. This pathway involves a serine protease cascade (Figure 2) that eventually cleaves an inactive protein called spa¨ tzle, making it competent to bind to a transmembrane receptor named Toll (Belvin and Anderson, 1996). Toll is homologous to the mammalian interleukin-1 receptor. Binding of spa¨ tzle to Toll initiates a signal transduction pathway that leads to activation of a transcription factor from the rel family named Dorsal. A large (350 kDa) multi-domain protein called nudel, containing a serine protease domain, regions of LDL receptor repeats (see Chapter 4.6), and an Nterminal glycosaminoglycan modification, is secreted

Figure 2 A model of the protease cascade that activates the Dorsal signal transduction pathway in D. melanogaster embryonic development. Nudel, gastrulation defective, snake, and easter are serine proteases that are synthesized as zymogens. The active forms of the proteases are indicated with an asterisk. Solid arrows indicate proteolytic activation steps that have been demonstrated by biochemical studies. A dotted arrow indicates that interaction between the snake and gastrulation defective zymogens can leads to activation of gastrulation defective. Easter* cleaves spa¨tzle to produce an active ligand that binds to Toll, a transmembrane receptor. Easter is negatively regulated by interaction with an inhibitor from the serpin family.

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by the ovarian follicle cells into the perivitelline space (Hong and Hashimoto, 1995; Turcotte and Hashimoto, 2002). The nudel protease is autoactivated by a mechanism not yet understood (LeMosy et al., 1998, 2000) and is thought to activate the second protease in the pathway, ‘‘gastrulation defective’’ by specific proteolysis. Mutations in nudel’s protease domain produce a dorsalizing phenotype and can also result in fragile eggshells, suggesting an additional function for the protease activity (Hong and Hashimoto, 1996; LeMosy et al., 1998, 2000; LeMosy and Hashimoto, 2000). Gastrulation defective is a serine protease (Konrad et al., 1998; Han et al., 2000), with sequence similarity to mammalian complement factors C2 and B (DeLotto, 2001). Experiments with recombinant proteins have demonstrated that the gastrulation defective zymogen can be autoactivated when it interacts with the protease snake zymogen and that active gastrulation defective can in turn proteolytically activate the snake zymogen (Dissing et al., 2001; LeMosy et al., 2001). Computer modeling studies indicate that the zymogen activation site of gastrulation defective is a good fit in the active site of nudel and that the snake zymogen activation site can dock in the active site of gastrulation defective, consistent with the proposed functions of these enzymes in the cascade pathway (Rose et al., 2003). A potential lower affinity interaction of the gastrulation defective active site with its own zymogen activation sequence may explain gastrulation defective’s autoactivation in the absence of nudel when it is overexpressed in embryos or at high concentration in vitro. The final two proteases in this cascade, snake and easter, contain C-terminal serine protease domains and N-terminal clip domains similar to horseshoe crab proclotting enzyme (DeLotto and Spierer, 1986; Chasan and Anderson, 1989; Gay and Keith, 1992; Smith and DeLotto, 1992, 1994). Clip domains, thought to function in protein–protein interactions, are also present in some hemolymph proteases of insects (Jiang and Kanost, 2000) (see Section 4.7.3.1 below). Mutations that eliminate the protease activity of snake (Smith et al., 1994) or easter (Jin and Anderson, 1990) have abnormal dorsoventral phenotypes, indicating that a functional protease domain is essential for their roles in embryonic development. In vitro experiments with recombinant snake and easter zymogens confirm their order in the cascade indicated by genetic analysis: snake cleaves and activates easter, which cleaves prospa¨tzle (Smith et al., 1995; DeLotto and DeLotto, 1998; Dissing et al., 2001; LeMosy et al., 2001). These results are consistent with predictions

250 Proteases

based on computer modeling of the snake and easter three-dimensional structures and substrate interactions sites (Rose et al., 2003). Active easter is converted in vivo to a high molecular mass form that is probably a complex with a protease inhibitor that regulates its activity (Misra et al., 1998; Chang and Morisato, 2002). Female flies with a mutation in the gene for a serine protease inhibitor, serpin 27A, produce embryos that show defects in dorsal– ventral polarity, suggesting that this inhibitor is a maternal product that regulates at least one of the proteases in the pathway (Hashimoto et al., 2003; Ligoxygakis et al., 2003).

4.7.3. Hemolymph Plasma Proteases 4.7.3.1. Serine Proteases p0060

Serine proteases in hemolymph have several types of physiological functions in defense against infection or wounding. An unusual phenomenon, perhaps related to protection against predation, involves serine proteases in the hemolymph of South American saturniid caterpillars of the genus Lonomia that are toxic to mammals. Contact with these caterpillars can result in acquired bleeding disorders due to potent fibrinolytic activity of these hemolymph proteases (Amarant et al., 1991; Arocha-Pinango and Guerrero, 2000). Extracellular serine protease cascades mediate rapid responses to infection and wounding in vertebrate and invertebrate animals. Biochemical and genetic evidence indicates that activation of serine proteases in arthropod hemolymph is a component of several immune responses, including coagulation, melanotic encapsulation, activation of antimicrobial peptide synthesis, and modulation of hemocyte function (Yoshida and Ashida, 1986; Katsumi et al., 1995; Kawabata et al., 1996; Ashida and Brey, 1998; Levashina et al., 1999; Jiang and Kanost, 2000; Gorman and Paskewitz, 2001; Kanost et al., 2001). Microbial challenge induces expression of many nondigestive serine proteases (Dimopoulos and Della Torre, 1996; Oduol et al., 2000; Irving et al., 2001; Dimopoulos et al., 2002; Zhu et al., 2003a). In Anopheles gamibiae, a large, multidomain serine protease, Sp22D, is expressed in hemocytes, fat body, and midgut and is secreted into the hemolymph, with a low level of induced expression in response to infection (Danielli et al., 2000; Gorman et al., 2000a). Sp22D contains chitinbinding domains, lipoprotein receptor class A domains, scavenger receptor domains, and a C-terminal serine protease domain. This complex domain architecture with multiple domains that

might function in binding to proteins or polysaccharides suggests that Sp22D may participate in the formation of protein complexes in defense responses or perhaps in tissue remodeling at metamorphosis. Most of the hemolymph proteases are expressed in fat body or hemocytes, but a bacteria-induced protease, scolexin, from Manduca sexta is expressed in epidermis (Kyriakides et al., 1995; Finnerty et al., 1999). Another novel serine protease expressed in pupal yellow body of Sarcophaga peregrina has antibacterial activity distinct from its protease activity (Nakajima et al., 1997; Tsuji et al., 1998). Serine proteases that contain a C-terminal protease domain and an N-terminal clip domain are known to act in cascade pathways in arthropod hemolymph (Jiang and Kanost, 2000). Among clip domain proteases with known function are horseshoe crab proclotting enzyme and clotting factor B (Kawabata et al., 1996), D. melanogaster snake and easter, and phenoloxidase-activating proteases described below. Clip domains are 35–55 amino acid residue sequences that contain three conserved disulfide bonds. They may function to mediate interaction of members of protease cascade pathways. Proteases may contain one or more N-terminal clip domains, followed by a 20–100 residue linking sequences connecting them to the catalytic protease domain. Insects that have been investigated in some detail are known to contain a large number of genes for clip domain proteases. Among the 204 genes with homology to the S1 serine protease family in the D. melanogaster genome, 24 are clip domain proteases, most of whose functions are unknown (Ross et al., 2003). The A. gambiae genome contains 41 clip domain proteases (Christophides et al., 2002) but only a few have been studied in detail (Han et al., 1997; Paskewitz et al., 1999; Gorman et al., 2000b; review: Gorman and Paskewitz, 2001). In M. sexta, more than 20 clip domain proteases expressed in fat body or hemocytes have been identified (Jiang et al., 1999; Jiang, Y. Wang and M.R. Kanost, unpublished results; Kanost et al., 2001). Melanization, a response to wounding and infection in insects and crustaceans, involves activation of a cascade of serine protease zymogens (Figure 3). This pathway leads to rapid activation of a protease that in turn activates a phenoloxidase zymogen (prophenoloxidase; proPO) (So¨ derha¨ll et al., 1996; Ashida and Brey, 1998). Oxidation of phenols by phenoloxidase leads to production of quinones that polymerize to form melanin (see Chapters 4.3 and 4.4). Melanization of encapsulated parasites is believed to be an important defensive response in insects, including insect vectors of human diseases

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Proteases

Figure 3 A model of the protease cascade that activates prophenoloxidase in response to infection. Hemolymph plasma proteins known as pattern recognition proteins bind to polysaccharides on the surface of microorganisms. This interaction leads to activation of initiator protease(s) by a mechanism not yet understood, which triggers a protease cascade. The number of proteases in the pathway is not yet known (indicated by dashed arrows). Activation of uncharacterized proteases leads to activation of activator of prophenoloxidase activating protease (PAP) and serine protease homologs (SPH) that function together to form a prophenoloxidase activating enzyme, which cleaves prophenoloxidase (ProPO) to form active phenoloxidase (PO). PO catalyzes the oxidation of hemolymph catecholic phenols to corresponding quinones, which can undergo further reactions to form melanin. Proteases in the pathway are regulated by serine protease inhibitors known as serpins.

(Gillespie et al., 1997; Paskewitz and Gorman, 1999). Serine proteases that activate prophenoloxidase have been purified, and corresponding cDNAs have been cloned from two lepidopteran insects: M. sexta (Jiang et al., 1998, 2003a, 2003b) and B. mori (Satoh et al., 1999); a beetle, Holotrichia diomphalia (Lee et al., 1998a, 1998b); and a crayfish (Wang et al., 2001). All of these enzymes contain a C-terminal serine protease catalytic domain and one or two N-terminal clip domains, similar to horseshoe crab clotting enzyme and D. melanogaster easter (Jiang and Kanost, 2000). These proteases are synthesized as zymogens, which must be activated by a protease upstream in the pathway. However, it is not known how many steps occur between recognition of a microorganism by a hemolymph protein and activation of the terminal protease in the pathway. Furthermore, it is not yet understood how the first protease in the cascade is activated. Proteins that bind to microbial surfaces (pattern recognition proteins) are probably involved in this step, through interaction with microbial polysaccharides and an initiating protease. Plasma proteins that bind to bacterial lipopolysaccharide,

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peptidoglycan, or fungal glucans can stimulate activation of the prophenoloxidase cascade (Ochiai and Ashida, 1988, 1999, 2000; So¨ derha¨ll et al., 1988; Yu et al., 1999, 2002; Ma and Kanost 2000; Yu and Kanost, 2000). Initial characterization of a proPO-activating proteinase in M. sexta indicated that the purified protease could not efficiently activate proPO, but required participation of a nonproteolytic protein fraction (Jiang et al., 1998). This protein cofactor was identified in H. diomphalia (Kwon et al., 2000) and M. sexta (Yu et al., 2003) as a protein with a clip domain and a serine protease domain, in which the active site serine residue is changed to glycine. There are 13 such clip domain serine protease homolog genes in the D. melanogaster genome (Ross et al., 2003). These serine protease homologs lack protease activity due to the incomplete catalytic triad and must therefore have other functions. One D. melanogaster serine protease homolog, masquerade, functions in nerve and muscle development (Morugasu-Oei et al., 1995, 1996). A serine protease homolog in crayfish hemolymph has a role in immune responses, indicating evolutionary conservation of function in these unusual proteins (Lee and So¨ derha¨ll, 2001). The active form of the serine protease homologs that function as cofactors for proPO activation are themselves activated through specific cleavage by a serine protease in hemolymph (Kim et al., 2002; Lee et al., 2002b; Yu et al., 2003) adding additional complexity to this pathway. The serine protease homologs from M. sexta that stimulate proPO activation bind to a hemolymph lectin, which is a recognition protein for lipopolysaccharides from Gram-negative bacteria, and to proPO and prophenoloxidase activating protease (Yu et al., 2003). The interaction between the lectin and a proPO activation complex may serve to localize melanin synthesis to the surface of invading bacteria. Serine proteases are also involved in other insect immune responses. The signal transduction system that regulates dorsal/ventral development in D. melanogaster embryos also regulates expression of the gene for an antifungal peptide in larvae and adults (review: Hoffmann, 2003). In embryonic development, this pathway involves an extracellular serine protease cascade that eventually cleaves an inactive protein, spa¨tzle, making it competent to bind to a transmembrane receptor named Toll. This same receptor–ligand interaction activates a signal pathway that leads to activation of rel family transcription factors that stimulate expression of drosomycin, an antifungal peptide normally synthesized by the fat body after microbial challenge.

252 Proteases

Mutants deficient in Toll or spa¨ tzle exhibit decreased induction of drosomycin (Lemaitre et al., 1996). These results suggest that infection leads to proteolytic processing of spa¨ tzle in the hemolymph. The activated spa¨tzle binds to the Toll receptor in fat body membranes, leading to activation of the Dif pathway and expression of drosomycin. Mutants of gastrulation defective, snake, or easter do not have an impaired antifungal response, indicating that a different set of proteases functions in the immune response protease cascade. Like prophenoloxidase activation, this pathway is initiated by interactions of pattern recognition proteins with microbial surface polysaccharides (Kim et al., 2000; Leulier et al., 2003), presumably stimulating activation of the first protease in the cascade. A clip domain protease known as persephone has been shown to participate in this pathway (Ligoxygakis et al., 2002a), but its position in the cascade is not yet known. 4.7.3.2. Protease Inhibitors p0100

Insect hemolymph contains high concentrations of serine protease inhibitors from several different gene families (reviews: Kanost and Jiang, 1996; Polanowski and Wilusz, 1996; Kanost, 1999). Protease cascade pathways in mammalian blood are regulated by 45 kDa protease inhibitors known as serpins (Silverman et al., 2001; Gettins, 2002). Serpins in arthropod hemolymph also function to regulate protease cascades, preventing detrimental effects of uncontrolled immune responses. For example, each of the proteases in the horseshoe crab coagulation pathway is regulated by serpins produced by hemocytes (Kawabata et al., 1996). Fourteen serpin genes have been identified in the A. gambiae genome (Christophides et al., 2002) and 26 serpin genes have been annotated in the D. melanogaster genome. In M. sexta, 7 serpin genes have been identified (Jiang et al., 1996; Gan et al., 2001; Zhu et al., 2003b). However, the physiological target proteases of only a few insect serpins have been determined (Ashida and Sasaki, 1994; Jiang et al., 2003b; Zhu et al., 2003a). The reactive site in a serpin protein that interacts with the target protease is part of an exposed loop near the C-terminal end of the serpin sequence. Some insect serpin genes have a unique structure in which mutually exclusive alternate splicing of an exon that encodes the reactive site loop results in production of several inhibitors with different specificity. This was first observed in the gene for M. sexta serpin-1, which contains 12 copies of its 9th exon. Each version of exon 9 encodes a different reactive

site loop sequence and inhibits a different spectrum of proteases (Jiang et al., 1996; Jiang and Kanost, 1997). Structures of two of the M. sexta serpin-1 variants have been determined by X-ray crystallography (Li et al., 1999; Ye et al., 2001). Serpin genes with alternate exons in the same position as in M. sexta serpin-1 have been identified in other insect species, including the lepidopterans B. mori (Sasaki, 1991; Narumi et al., 1993) and Mamestra configurata (Chamankhah et al., 2003), in the dipterans D. melanogaster (Kruger et al., 2002) and A. gambiae (Danielli et al., 2003), and in the cat flea, Ctenocephalides felis (Brandt et al., 2004). Like blood clotting, phenoloxidase activation is normally regulated in vivo as a local reaction of brief duration. This regulation involves serine protease inhibitors in plasma (Kanost and Jiang, 1996; Kanost, 1999). Two serpins from the hemolymph of M. sexta (serpin-1J and serpin-3) can inhibit proPO activating proteases (Jiang et al., 2003a, 2003b; Zhu et al., 2003a, 2003b). In D. melanogaster, serpin 27A (an apparent ortholog of Manduca serpin-3) is involved in regulation of melanization (DeGregorio et al., 2002; Ligoxygakis et al., 2002b). Similarly, a mutation in D. melanogaster serpin 43Ac (Necrotic) leads to constitutive expression of drosomycin, indicating that this serpin regulates a protease in the cascade that processes spa¨ tzle (Figure 4) (Levashina et al., 1999; Green et al., 2000). It is not yet known whether serpin 43Ac inhibits persephone or some other protease in the pathway. In addition to serpins, lower molecular weight inhibitors from the Kunitz family (Sugumaran et al., 1985; Saul and Sugumaran, 1986; Aso et al., 1994) and a family of 4 kDa inhibitors from locusts (Boigegrain et al., 1992) can interfere with proPO activation (review: Kanost, 1999), although it is not yet known which proteases in the pathway they can inhibit.

4.7.4. Cellular Proteases 4.7.4.1. Serine Proteases

Serine proteases appear to function in tissue remodeling and extracellular matrix degradation required for cell movements in metamorphosis. Evagination of D. melanogaster imaginal discs is blocked by serine protease inhibitors, and the discs release serine proteases (Pino-Heiss and Schubiger, 1989). The Stubble-stubbloid gene encodes an integral membrane protein with an extracellular serine protease domain. It is expressed in imaginal discs under control of 20-hydroxyecdysone (see

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Natori, 1996). A similar cathepsin L from another dipteran, Delia radicum, is expressed highly in midgut beginning in late third instar and may function in metamorphosis of the midgut (Hegedus et al., 2002). A cathepsin L-like protease expressed in a D. melanogaster haemocyte cell line is present in granules and may be a lysosomal enzyme functioning to degrade phagocytosed material (Tryselius and Hultmark, 1997). 4.7.4.3. Caspases

Figure 4 A model of the protease cascade that activates the D. melanogaster Toll pathway for synthesis of antimicrobial peptides. Hemolymph plasma proteins known as pattern recognition proteins bind to polysaccharides on the surface of microorganisms. This interaction leads to activation of initiator protease(s) by a mechanism not yet understood, which triggers a protease cascade. The number of proteases in the pathway is not yet known (indicated by dashed arrows). A clip domain protease called Persephone is known from genetic data to be a component in the pathway. The cascade leads to activation of a protease (designated as X in this figure) that cleaves spa¨tzle to produce an active ligand that binds to Toll, a transmembrane receptor. A serpin, Spn43Ac negatively regulates at least one protease in the pathway.

Chapter 3.3) and may function to detach imaginal discs from extracellular matrices (Appel et al., 1993). 4.7.4.2. Cathepsin-Type Cysteine Proteases

Cysteine proteases related to cathepsin B and cathepsin L have been identified as proteins produced by hemocytes that participate in tissue remodeling in metamorphosis of several insects. In S. peregrina, a 26/29 kDa protease synthesized in hemocytes was identified as a cathepsin B (Saito et al., 1992; Kurata et al., 1992a; Takahashi et al., 1993; Fujimoto et al., 1999). This protease may be released from pupal hemocytes to cause dissociation of fat body at metamorphosis (Kurata et al., 1992b). A cathepsin B from hemocytes of B. mori may also function in tissue degradation during metamorphosis, including histolysis of silk glands (Shiba et al., 2001; Xu and Kawasaki, 2001; Chapter 2.11). Cysteine proteases classified as cathepsin L have also been identified as participants in tissue remodeling at metamorphosis. A cathepsin L from S. peregrina appears to function in the differentiation of imaginal discs (Homma et al., 1994; Homma and

Programmed cell death, known as apoptosis, is an essential process in development (Richardson and Kumar, 2002; Chapter 2.5) and is also a response by insect cells to viral infection (Clarke and Clem, 2003). An intracellular cascade of cysteine proteases called caspases (MEROPS family C14) is a key pathway in initiating apoptosis. Caspases are conserved in structure and function in animal species. They cleave target protein substrates after Asp or Glu P1 residues. Caspases are synthesized as inactive zymogens. They are activated by specific proteolytic cleavage, yielding a small and a large subunit that together form an active site. These heterodimers further dimerize to form a hetrotetramer with two active sites (Earnshaw et al., 1999). Caspases are divided into two groups, initiator caspases that interact with apoptosis-initiating proteins, and effector caspases, which are activated by the initiators. Initiator caspases contain a C-terminal cysteine protease domain and a long N-terminal region that contains death effector domains (DEDs) or caspase recruitment domains (CARDs), which interact with proteins that stimulate apoptosis. Dimerization of initiator caspases occurs when they associate with apoptosis-promoting proteins, causing caspase activation through conformational change in the absence of proteolytic cleavage (Boatright et al., 2003). The initiator caspases then activate the effector caspases through specific proteolytic cleavage. Effector caspases, which lack a long N-terminal domain, cleave structural components of the cytoskeleton and nucleus and proteins involved in signaling pathways, resulting in death of the cell. Cleavage by effector caspases activivates death-promoting enzymes, including certain kinases and nucleases (Earnshaw et al., 1999). Apoptosis and caspases have been investigated in detail in D. melanogaster (Figure 5). The D. melanogaster genome contains seven caspase genes. Three genes encode caspases with long prodomains (dronc, dcp-2/dredd, and strica/dream) and four genes encode caspases with short prodomains (dcp-1, drICE, decay, and damm/daydream) (Vernooy et al., 2000; Kumar and Doumanis, 2000).

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Figure 5 A model for a caspase cascade pathway in apoptosis in D. melanogaster. Cellular proapoptotic protein factors initiate oligomerization of the caspase zymogen proDRONC, which results in its self-processing and activation. Active DRONC cleaves the effector caspases DrICE and DcP-1, which then become active and cleave protein substrates including inactive forms of nucleases and kinases. The resulting active enzymes then catalyze reactions that result in progressive disassembly of the nucleus and cytoskeleton and cell death. A caspase inhibitor DIAP1 inhibits DRONC, DCP-1, and DrICE to regulate their activities.

Dronc contains an N-terminal CARD, and its overexpression in flies or in cultured cells stimulates apoptosis (Dorstyn et al., 1999a; Quinn et al., 2000). Dronc expression is stimulated by ecdysteroid, and it appears to function in apoptosis in tissue remodeling at metamorphosis (Dorstyn et al., 1999a; Cakouros et al., 2002; Lee et al., 2002b; Chapter 2.5). Depletion of Dronc by RNA interference results in a decrease in normal programmed cell death in embryogenesis (Quinn et al., 2000). Dronc can cleave substrates with P1 Glu or Asp residues. It can autoactivate by cleaving following a glutamate residue, but it activates drICE by cutting at an aspartate residue (Hawkins et al., 2000). Dredd is an initiator caspase that participates in apoptosis (Chen et al., 1998). Dredd also functions in a signal transduction pathway that leads to activation of immune responses (Elrod-Erickson et al., 2000; Leulier et al., 2000; Georgel et al., 2001) through its cleavage and activation of a rel family transcription factor called relish (Stoven et al., 2000, 2003). The third initiator caspase in D. melanogaster, STRICA, lacks either a CARD or a DED and instead contains a serine/threonine rich prodomain, suggesting that STRICA functions in an as yet unidentified pathway (Doumanis et al., 2001). The effector caspases Dcp-1 and DrICE are processed by active DRONC and then cleave target proteins that result in the disassembly of the cell.

Both caspases induce apoptosis when overexpressed, have preferred target cleavage sites very similar to those of mammalian caspase-3, and can be inhibited by the baculovirus protein P35, a powerful inhibitor of effector caspases (Fraser and Evan, 1997; Fraser et al., 1997; Song et al., 1997; Meier et al., 2000). Dcp-1 and DrICE have slight differences in specificity. Active DCP-1 can activate other proDCP-1 proteins and proDrICE, whereas active DrICE can not cleave and activate proDrICE (Song et al., 2000). A third effector caspase, DECAY, shares similarity with DrICE and DCP-1 in sequence and preferred cleavage site, and can induce apoptosis when ectopically expressed in cultured cells (Dorstyn et al., 1999b). In contrast, the shortprodomain caspase DAMM appears to have an accessory role in apoptosis, sensitizing the cell to prodeath signals rather then directly killing the cell. DAMM shares strong sequence similarity with STRICA, has a preferred cleavage site similar to mammalian caspases -1, -4, and -5, and is not processed in vitro by DAMM, DRONC, DECAY, DCP-1, or DrICE. DAMM mRNA is upregulated in apoptotic tissues, and DAMM over-expression in eye tissue sensitizes the cells to apoptosis from either developmental signals or radiation (Harvey et al., 2001). The caspase pathways in other insects may be even more complex. For example, the A. gambiae genome contains 12 caspase genes, compared to 7 in D. melanogaster. An effector caspase has been identified in the lepidopteran Spodptera frugiperda (Ahmad et al., 1997; Seshagiri and Miller, 1997). This enzyme, Sf-caspase-1, is activated by an initiator caspase equivalent to DRONC (Manji and Friesen, 2001). The three-dimensional structure of Sf-caspase-1 has been determined (the first structure of a nonhuman caspase) by X-ray crystallography (Forsyth et al., 2004) in the presence of a tetrapeptide inhibitor. The Sf-caspase-1 fold is very similar to that of human effector caspases, but it has unique features in its N-terminal prodomain that may be important in regulating its activation. Caspases are regulated by specific inhibitor proteins that are present in the cytoplasm. In D. melanogaster, cells appear to be constitutively primed for caspase activation, and removal of caspase inhibition is sufficient to cause cell death. Three inhibitors of apoptosis (IAP) proteins, which inhibit caspases, have been identified in D. melanogaster (Hay et al., 1995; Jones et al., 2000). DIAP1 inhibits Dronc, DrICE, and DCP-1 (Kaiser et al., 1998; Hawkins et al., 1999; Meier et al., 2000; Wilson et al., 2002). D. melanogaster exhibits a constitutive proapoptotic predisposition, resulting in rapid apoptosis of cultured cells or embryos in the event of

Proteases

the decreased expression of DIAP1 (Igaki et al., 2002; Muro et al., 2002; Rodriguez et al., 2002). Iap genes, similar to D. melanogaster diap1 and sharing the same capability of inhibiting initiator caspases, have been identified from three lepidopterans, Trichoplusia ni (Liao et al., 2002; Seshagiri et al., 1999), B. mori (Huang et al., 2001), and S. frugiperda (Huang et al., 2000). RNA interference depletion of Sf-IAP from S. frugiperda cell lines results in apoptosis, suggesting that, like Drosophila cells, Spodoptera cells are maintained in a state predisposed to cell death (Muro et al., 2002). Baculoviruses have evolved two types of caspase inhibitors that can block the lepidopteran apoptotic program. AcMNPV expresses its gene for the antiapoptotic protein P35 at both early and late times during an infection, thus preventing infected cells from prematurely terminating viral replication through apoptosis (Clem et al., 1991). P35 is a globular protein with an extended loop (Fisher et al., 1999; Zoog et al., 1999) that contains a caspase cleavage site recognized by a wide range of effector caspases, including human caspases-1, -3, -6, -7, -8, -10 (Zhou et al., 1998), Sf-caspase-1 (Ahmad et al., 1997), and DrICE (Fraser et al., 1997), but not by initiator caspases such as human caspase-9 (Vier et al., 2000), DRONC (Meier et al., 2000), or Sf-caspase-X (LaCount et al., 2000). When an effector caspase cleaves in the reactive site loop of P35, a thioester bond forms between the active site cysteine of the caspase and the P1 aspartic acid residue of P35, causing a conformational change in P35 that blocks access to water molecules, thus preventing completion of the peptide bond hydrolysis (Bertin et al., 1996; dela Cruz et al., 2001; Riedl et al., 2001; Xu et al., 2001). Baculoviruses also contain genes for IAP proteins that block apoptosis by inhibiting initiator caspases (Huang et al., 2000b; Birnbaum et al., 1994; LaCount et al., 2000; Means et al., 2003) suggesting that, at some point during their evolution, baculoviruses co-opted the cell’s own mechanism for regulating caspase activity. In fact, the first IAP protein to be discovered was Cp-IAP from the baculovirus Cydia pomonella granulovirus (Crook et al., 1993). 4.7.4.4. Metalloproteases

Three families of metalloproteases that are not involved in digestion of food have been targets of fairly limited investigation in insects. They appear to function in remodeling of the extracellular matrix or in degradation of peptide hormones. Matrix metalloproteases (MMP) are integral membrane proteins, present on the outer surface of

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cells. They are multidomain proteins that include a catalytic domain that incorporates a zinc ion in the active site. In mammals, these enzymes regulate processes involving morphogenesis in development and tissue remodeling by digesting protein components of the extracellular matrix (Nagase and Woessner, 1999). In D. melanogaster there are two MPP genes. Dm1-MMP is expressed strongly in embryos and may have a role in remodeling of the extracellular matrix in development of the central nervous system (Llano et al., 2000). Dm2-MMP is expressed at a low level at all developmental stages, but with strong expression in regions of the brain and the eye imaginal discs (Llano et al., 2002). Mutants in Dm1-MMP have defects in larval tracheal development and pupal head eversion, whereas mutants in Dm2-MMP do not undergo proper tissue remodeling during metamorphosis. However, normal embryonic development was observed even in double mutants lacking both MMPs (PageMcCaw et al., 2003). It appears that these proteases are required primarily for remodeling of the extracellular matrix in metamorphosis. In mammals, the MMPs are regulated by proteins called tissue inhibitors of metalloproteases (TIMP). A homolog of these proteins in D. melanogaster has been shown to inhibit its MMPs (Wei et al., 2003). A different class of metalloproteases known as ADAMs (because they contain a disintegrin and metalloprotease domain) are also integral membrane proteins, with the zinc-containing protease domain on the extracellular surface. In mammals, they participate in growth factor processing, cell adhesion, cell fusion, and tissue remodeling processes, although their physiological functions, particularly of the protease domain, are still not well understood (Seals and Courtneidge, 2003). A D. melanogaster ADAM called kuzbanian has been shown to function in axon extension in nervous system development (Fambrough et al., 1996; Rooke et al., 1996). Kuzbanian is involved in initiation of a signal transduction pathway by a transmembrane receptor called Notch (Sotillos et al., 1997). The metalloprotease domain of Kuzbanian may exert its physiological effect through cleavage of Notch (Lieber et al., 2002) or the Notch ligand, Delta (Bland et al., 2003). A third family of zinc metalloproteases identified in insects contains members that may function in the degradation of peptide hormones. These transmembrane proteins are similar to mammalian neprilysins (also called neutral endopeptidases), which cleave oligopeptides on the amino side of hydrophobic residues. They cleave physiologically active signaling peptides with functions in nervous, cardiovascular,

256 Proteases

inflammatory, and immune systems and have a wide tissue distribution (Turner et al., 2001). Metalloprotease activities with properties similar to neprylins have been identified as enzymes that can degrade tachykinin-related peptides in a cockroach, Leucophaea maderae, a locust, Locusta migratoria, a dipteran, D. melanogaster, and a lepidopteran, Lacanobia oleracea (Isaac et al., 2002; Isaac and Nassel, 2003). Similar activities that degrade adipokinetic hormone have been identified in the lepidopterans Lymantria dispar (Masler et al., 1996) and M. sexta (Fox and Reynolds, 1991), and a dipteran, M. domestica (Lamango and Isaac, 1993). cDNA clones for proteases with sequence similarity to neprilysin have been described in B. mori (Zhao et al., 2001), M. sexta (Zhu et al., 2003a), and L. migratoria (Macours et al., 2003). The D. melanogaster genome contains 24 neprilysin-like genes (Coates et al., 2000). Further study is needed to determine the substrates and physiological roles of these proteases, but it seems likely that they function as negative regulators of peptide hormones. 4.7.4.5. Aspartic Acid Proteases

The aspartic acid proteases from the MEROPS families A1 and A2 (similar to human pepsin) are a group that has received little study in insects. The D. melanogaster genome contains 34 genes that encode proteins similar to these enzymes, but none appear to have been investigated at the molecular level. A cathepsin D-like aspartic acid protease from A. aegypti has been identified as a lysosomal enzyme, which accumulates in fat body during vitellogenin synthesis (Cho and Raikhel, 1992; Dittmer and Raikhel, 1997). An enzymatically inactive protein from the A1 family is an important allergen from a cockroach (Blattella germanica) that triggers asthmatic responses in humans (Pome´ s et al., 2002). It is present at highest concentration in the gut (Arruda et al., 1995), but since it apparently lacks proteolytic activity, its function in the insect is unknown. 4.7.4.6. Proteasomes

Proteasomes are complex intracellular proteases that function in regulated degradation of cellular proteins. Turnover of proteins by the proteasome regulates many processes including the cell cycle, circadian cycles (see Chapter 4.11), transcription, growth, development, as well as removal of abnormal proteins. Proteins are targeted for degradation by the proteasome by attachment of polyubiquitin chains to an amino group on a lysine side chain. Eukaryotic proteasomes are composed of four

stacked heptameric rings that form a cylinder with multiple protease catalytic sites in its interior. This structure, the 20S proteasome, is composed of 28 subunits from multiple homologous gene products and has a mass of ~700 kDa. The 20S proteasome has little activity unless it is activated by another 700 kDa, 20 subunit protein, PA700, that can bind to one or both ends of the cylinder. When both ends of the 20S proteasome are capped by a PA700, the resulting complex is the 26S proteasome, which is active in degrading ubiquitinated proteins, an ATPdependent process (DeMartino and Slaughter, 1999). Proteasomes in insects have been studied primarily in D. melanogaster and M. sexta (Mykles, 1997, 1999). They have physical and catalytic properties similar to those of proteasomes from other eukaryotic species (Uvardy, 1993; Haire et al., 1995; Walz et al., 1998). Mutations in genes for subunits of D. melanogaster proteasomes or proteins, that regulate proteasome activity, result in lethal or otherwise complex phenotypes involving the disruption of multiple aspects of physiology and development (Schweisguth, 1999; Ma et al., 2002; Watts et al., 2003). Some mutations alter subunit composition or disrupt proteasome assembly (Covi et al., 1999; Smyth and Belote, 1999; Szlanka et al., 2003). Recent studies have used double-stranded RNA interference to study disruptions caused by reduced expression of individual proteasome components (Wojcik and DeMartino, 2002; Lundgren et al., 2003). In M. sexta, programmed cell death of intersegmental muscles at metamorophosis is accompanied by marked changes in proteasome activity and subunit composition (Dawson et al., 1995; Jones et al., 1995; Takayanagi et al., 1996; Low et al., 1997, 2000, 2001; Hastings et al., 1999).

4.7.5. Conclusions and Future Prospects Tremendous advances have been made in the last 10 years in our knowledge of the existence, structure, and function of insect proteases that have biological roles unrelated to digestion of food. Much more remain to be discovered. Through examination of the genome sequences of D. melanogaster and A. gambiae, we can see that these insects have an enormous number of genes encoding proteases and that most of them are unstudied and have unknown functions. Nearly all of the research in this area has focused on a few dipteran and lepidopteran species. It is to be expected that detailed investigation of a broader range of species will reveal complex and diverse functions for proteases

Proteases

in regulating intracellular and extracellular processes. A common feature of proteases is that they are synthesized as inactive zymogens, activated by proteolysis when the time is right, and then rapidly inhibited by specific inhibitors. Better understanding of molecular mechanisms of this tight regulation of multiple and varied protease cascade pathways will impact many areas of insect biology.

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challenge, and malaria infection. Proc. Natl Acad. Sci. USA 99, 8814–8819. Dimopoulos, G., Richman, A., della Torre, A., Kafatos, F.C., Louisn, C., 1996. Identification and characterization of differentially expressed cDNAs of the vector mosquito, Anopheles gambiae. Proc. Natl Acad. Sci. USA 93, 13066–13071. Dissing, M., Giordano, H., DeLotto, R., 2001. Autoproteolysis and feedback in a protease cascade directing Drosophila dorsal–ventral cell fate. EMBO J. 20, 2387–2393. Dittmer, N.T., Raikhel, A.S., 1997. Analysis of the mosquito lysosomal aspartic protease gene: an insect housekeeping gene with fat body-enhanced expression. Insect Biochem. Mol. Biol. 27, 323–335. Dorstyn, L., Colussi, P.A., Quinn, L.M., Richardson, H., Kumar, S., 1999a. DRONC, an ecdysone-inducible Drosophila caspase. Proc. Natl Acad. Sci. USA 96, 4307–4312. Dorstyn, L., Read, S.H., Quinn, L.M., Richardson, H., Kumar, S., 1999b. DECAY, a novel Drosophila caspase related to mammalian caspase-3 and caspase-7. J. Biol. Chem. 274, 30778–30783. Doumanis, J., Quinn, L., Richardson, H., Kumar, S., 2001. STRICA, a novel Drosophila melanogaster caspase with an unusual serine/threonine-rich prodomain, interacts with DIAP1 and DIAP2. Cell. Death Differ. 8, 387–394. Earnshaw, W.C., Martins, L.M., Kaufmann, S.H., 1999. Mammalian caspases: structure, activation, substrates, and functions during apoptosis. Annu. Rev. Biochem. 68, 383–424. Elrod-Erickson, M., Mishra, S., Schneider, D., 2000. Interactions between the cellular and humoral immune responses in Drosophila. Curr. Biol. 10, 781–784. Fambrough, D., Pan, D., Rubin, G.M., Goodman, C.S., 1996. The cell surface metalloprotease/disintegrin Kuzbanian is required for axonal extension in Drosophila. Proc. Natl Acad. Sci. USA 93, 13233–13238. Felsted, R.L., Kramer, K.J., Law, J.H., Berger, E., Kafatos, F.C., 1973. Cocoonase IV. Mechanism of activation of prococoonase from Antheraea polyphemus. J. Biol. Chem. 248, 3021–3028. Finnerty, C.M., Karplus, P.A., Granados, R.R., 1999. The insect immune protein scolexin is a novel serine proteinase homolog. Protein Sci. 8, 242–248. Fisher, A.J., Cruz, W., Zoog, S.J., Schneider, C.L., Friesen, P.D., 1999. Crystal structure of baculovirus P35: role of a novel reactive site loop in apoptotic caspase inhibition. EMBO J. 18, 2031–2039. Forsyth, C.M., Lemongello, D., LaCount, D.J., Friesen, P.D., Fisher, A.J., 2004. Crystal structure of an invertebrate caspase. J. Biol. Chem. 279, 7001–7008. Fox, A.M., Reynolds, S.E., 1991. Degradation of adipokinetic hormone family peptides by a circulating endopeptidase in the insect Manduca sexta. Peptides 12, 937–944. Fraser, A.G., Evan, G.I., 1997. Identification of a Drosophila melanogaster ICE/CED-3-related protease, drICE. EMBO J. 16, 2805–2813.

Proteases

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4.8 Lipocalins and Structurally Related Ligand-Binding Proteins H Kayser, University of Ulm, Germany ß 2005, Elsevier BV. All Rights Reserved.

4.8.1. Introduction 4.8.2. The b-Barrel Structural Motif 4.8.3. Biliproteins: Prototypic Lipocalins 4.8.3.1. Biliproteins with Known Crystal Structure 4.8.3.2. Biliproteins with Unknown Crystal Structure 4.8.3.3. Molecular Variants of Biliproteins 4.8.3.4. Developmental Expression and Biosynthesis of Biliproteins 4.8.3.5. Putative Functions of Biliproteins 4.8.4. Nitrophorins and Related Proteins 4.8.4.1. The Problem of Blood Feeders 4.8.4.2. Crystal Structures of Nitrophorins from Rhodnius prolixus 4.8.4.3. Other Lipocalins from Blood-Feeding Insects 4.8.4.4. Lipocalins from Blood-Feeding Ticks 4.8.4.5. Developmental Expression of Nitrophorins in Rhodnius prolixus 4.8.4.6. Functions of Nitrophorins and Related Proteins 4.8.5. Lipocalins with Unknown Ligands 4.8.5.1. Lipocalins Putatively Related to Development 4.8.5.2. Lipocalins Putatively Related to Reproduction 4.8.6. Fatty Acid-Binding Proteins 4.8.6.1. Variation of the Lipocalin Structure 4.8.6.2. Fatty Acid-Binding Proteins with Known Crystal Structure 4.8.6.3. Fatty Acid-Binding Proteins with Unknown Crystal Structure 4.8.6.4. Functions and Developmental Expression of Fatty Acid-Binding Proteins 4.8.7. Lipocalins and Fatty Acid-Binding Proteins as Allergens 4.8.8. Ligand-Binding Proteins That Are Not b-Barrels 4.8.8.1. Odorant-Binding Proteins 4.8.8.2. Lipophorins 4.8.9. Crustacean Lipocalins with Established Crystal Structure 4.8.10. Lipocalin Engineering: An Insect Protein Takes the Lead 4.8.11. Lipocalins – One Fold, Many Roles: Summary and Outlook

4.8.1. Introduction Studies of insects have significantly contributed to the establishment of the lipocalins as a new family of proteins. Members of the lipocalin family are defined by a specific three-dimensional motif of polypeptide folding and by the binding of small lipophilic molecules in their interior. Starting about 20 years ago, the number of lipocalins has increased enormously through the structural analysis of known proteins as well as the discovery of new proteins showing the lipocalin structural motif. Today, the lipocalins represent a superfamily of proteins including members from vertebrates, insects, mites, ticks, crustaceans, other invertebrates, and, more recently, from plants and even prokaryotes. This wide occurrence of lipocalins is thought to reflect a common ancestry and a long evolutionary history

267 268 270 270 274 275 276 278 279 279 280 282 282 285 285 286 286 289 289 289 290 293 293 294 295 295 295 296 297 299

suggesting also the involvement of these proteins in many vital processes (Ganfornina et al., 2000). The relatively late discovery of the lipocalins as a structural family of proteins is likely due to a low level of homology of their amino acid sequences. In pairwise comparisons of sequences, the identity between lipocalins may be below 20%, which is regarded as the lowest level for reliable alignments. Alternatively, lipocalins share a few sequence motifs in structurally conserved regions besides their overall highly conserved three-dimensional structure. Since sequence homology is of only limited value in identifying a new protein as a lipocalin, crystal structures have to be established for a definite assignment to this family, if modeling of its sequence into a known lipocalin structure is not successful. Hence, progress in the molecular characterization of

268 Lipocalins and Structurally Related Ligand-Binding Proteins

lipocalins depends on the availability of the protein of interest in sufficient quantity obtained either from a biological source or by heterologous expression of its cDNA. It depends further on the feasibility of obtaining well-diffracting crystals and, of course, on technical advancement in the crystallography field. The rapid growth of knowledge of the lipocalins with respect to structure and function in the past decade is accounted for by a recent special edition of Biochimica Biophysica Acta (vol. 1482, nos. 1–2, 2000) devoted entirely to this family of proteins. This issue offers a collection of relatively recent overviews of the lipocalin field dealing with general aspects of structures and functions as well as individual lipocalins including some from insects. The progress and growing importance of the lipocalins is further demonstrated by the first international Lipocalin Conference, which was held in 2003 in Copenhagen (Denmark). Those readers interested in the world of lipocalins are also referred to the lipocalin website. The significant contributions from insect studies to the discovery and understanding of the lipocalins have not yet been comprehensively documented. The intention, therefore, of this chapter is to present an overview of the lipocalins identified in insects to date, including the fatty acid-binding proteins as structurally closely related ligand-binding proteins. Since all these diverse proteins represent variations of a common core structural theme, called an antiparallel b-barrel, this chapter describes the group-specific polypeptide folding and the underlying primary structures followed by the biological context, which comprises the occurrence of these proteins and the expression patterns of the corresponding genes, as well as the established and putative functions of each type of protein. The few lipocalins and structurally related proteins from ticks, mites, and crustaceans are included to cover all arthropods. Two well-known families of insect ligand-binding proteins that turned out not to be constructed as b-barrels, however, are also described to demonstrate their alternate folding. The lipocalins and fatty acid-binding proteins referred to in this chapter are listed in Table 1, including their accession codes. All multiple alignments of protein sequences shown in this chapter were performed using Clustal W, version 1.82, accessed via ExPASy. Graphical representations of crystal structures were prepared with Programme O (Jones et al., 1991).

4.8.2. The b-Barrel Structural Motif The characteristic lipocalin fold was first described for the human retinol-binding protein 20 years ago

(Newcomer et al., 1984), which thus became a kind of reference or prototypic lipocalin. Retinol-binding protein was soon followed by another vertebrate protein, b-lactoglobulin, the crystal structure of which showed high similarity to that of retinolbinding protein, though their sequences were not closely related (Papiz et al., 1986). At this very early stage of the analysis of the lipocalin family, two insect proteins were studied independently for their crystal structures and, again, the close similarities in the folding motifs to that of retinol-binding protein were recognized instantly (Holden et al., 1987; Huber et al., 1987a, 1987b). These insect proteins were two lepidopteran biliproteins, which are discussed in detail below (see Section 4.8.3.1). The discovery of the architectural principle of retinol-binding protein in two evolutionarily distant insect proteins immediately suggested this new folding type to be very common across the phyla, though obviously not restricted to a specific functional class of proteins. Typically, lipocalins are small extracellular, i.e., secreted, proteins of a size in the 20 kDa range, corresponding to about 180 amino acid residues, with affinity for specific small lipophilic molecules. Therefore, lipocalins are often referred to as extracellular lipid-binding proteins (eLBPs) to differentiate them from the related intracellular type (iLBPs) (see Section 4.8.6). The remarkably conserved lipocalin tertiary protein structure is called a b-barrel with a repeated þ1 topology. This means it is made up of eight sequentially arranged antiparallel b-strands, assigned A to H, which are hydrogenbonded to form a b-sheet that is folded back on itself to form an orthogonal stacking of two layers. The result is a barrel-like structure of a spherical shape. The eight b-strands are linked by seven loops, L1 to L7, which are all very short b-hairpins, except L1 that forms a large O loop covering partially the barrel opening with the internal ligand binding site. The other end of the barrel is closed by an N-terminal short 310-helical structure, which is followed immediately by strand A. One side of the barrel is lined with the C-terminal sequence, which includes a conserved a-helix preceding the short extra b-strand I, which does not contribute to the barrel structure. There are typically three conserved motifs, called structurally conserved regions (SCRs), each of them being characterized by a more or less extended invariant sequence motif. Two of the conserved regions, SCR1 and SCR3, which are present in all lipocalins, are located within the N- and C-terminal sequences comprising the 310-helix þ strand A and strand H þ the loop to the C-terminal a-helix, respectively. Another region, SCR2, that may be less

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269

Table 1 Arthropod lipocalins and fatty acid-/retinoic acid-binding proteins referred to in the text, and their accession codes

Name of protein

Source

Sequence (SwissProt/ TrEMBL)

Pieris brassicae Manduca sexta Manduca sexta Manduca sexta Samia cynthia ricini Samia cynthia ricini Rhodnius prolixus Rhodnius prolixus Rhodnius prolixus Rhodnius prolixus Triatoma pallidipennis Triatoma pallidipennis Rhodnius prolixus Rhodnius prolixus Rhodnius prolixus Rhipicephalus appendiculatus

P09464 P00305 Q00629 Q00630 Q8T118 Q8T119 Q26239 Q26241 Q94733 Q94734 Q27049 Q27042 Q94731 Q94732 Q86PT9 O77420

1BBP coord. not in data bank

Rhipicephalus appendiculatus

O77421

1QFT, 1QFV

Rhipicephalus appendiculatus

O77422

Dermacentor reticulatus Triatoma protracta Blattella germanica Schistocerca americana Drosophila melanogaster Drosophila melanogaster Galleria mellonella Bombyx mori Hyphantria cunea Leucophaea maderae Leucophaea maderae Homarus gammarus Homarus gammarus Homarus gammarus Homarus gammarus

Q8WSK7 Q9U6R6 P54962 P49291 Q9NAZ3 (Q9V6K5) Q9NAZ4 (Q8SXR1) Q24996 Q95P97 Q8T5Q9 O46130 Q95VP9 P58989

Manduca sexta Manduca sexta Schistocerca gregaria Locusta migratoria Heliothis zea Drosophila melanogaster Anopheles gambiae Apis mellifera Blomia tropicalis Lepidoglyphus destructor Acarus siro Manduca sexta

P31416 P31417 P41496 P41509 O76515 Q9VGM2 Q17017 Q9Y1C6 Q17284 Q9U5P1 O76821 O61236

Crystal structure (PDB)

Lipocalins

Bilin-binding protein Insecticyanin a, mature form Insecticyanin a Insecticyanin b Biliverdin-binding protein I Biliverdin-binding protein II Nitrophorin 1 Nitrophorin 2 Nitrophorin 3 Nitrophorin 4 Triabin Pallidipin 2 Salivary platelet aggregation inhibitor 1 Salivary platelet aggregation inhibitor 2 Biogenic amine-binding protein Histamine-binding protein 1, female specific Histamine-binding protein 2, female specific Histamine-binding protein 3, male specific Serotonin-histamine-binding protein Procalin Bla g 4 Lazarillo glial Lazarillo neural Lazarillo Gallerin Bombyrin Hyphantrin Tergal gland protein Lma-P22 Tergal calycin p18 Crustacyanin A1 subunit Crustacyanin A1 subunit Crustacyanin A2 subunit Crustacyanin C1 subunit

P80007 P80029

1NP1; 2NP1; 3NP1; 4NP1 1EUO 1NP4; 1ERX; 1IKE; 1KOI; 1IKJ 1AVG

1GKA (holoA1/A2-dimer) 1H91 (apoA1/A1-dimer) 1I4U (apoC1/C1-dimer)

Fatty acid-/retinoic acid-binding proteins

Fatty acid-binding protein MFB1 Fatty acid-binding protein MFB2 Fatty acid-binding protein Fatty acid-binding protein Fatty acid-binding protein Fatty acid-binding protein Fatty acid-binding protein Fatty acid-binding protein Fatty acid-binding protein Blo t 13 Fatty acid-binding protein Lep d 13 Fatty acid-binding protein Aca s 13 Cellular retinoic acid-binding protein

1MDC 1FTP

PDB, protein data bank.

well conserved, covers parts of the strands F and G and the connecting loop L6 at the bottom of the barrel. All SCRs are located at the closed end of the b-barrel which may provide a docking site for other

proteins. Furthermore, lipocalin sequences display typically four cysteine residues that form two disulfide bridges at comparable positions. The typical arrangement of secondary structures and structurally

270 Lipocalins and Structurally Related Ligand-Binding Proteins

Figure 1 Alignment of the biliproteins with known full sequences. The proteins are insecticyanin a (INSa) and insecticyanin b (INSb) from Manduca sexta, the bilin-binding protein (BBP) from Pieris brassicae, and the biliverdin-binding proteins I and II (SAMI and SAMII) from Samia cynthia ricini. Black boxes show residues typically conserved in lipocalins. N-terminal secretion signals are boxed in gray and the structurally conserved regions SCR1, SCR2, and SCR3 are in yellow, green, and orange, respectively. Below the sequences, the approximate positions of helices and b-strands are labeled as red and blue bars, respectively. Asterisks indicate identical residues, double and single points more or less conserved substitutions.

conserved regions is illustrated in Figure 1 with sequences from biliproteins representing prototypic lipocalins (see Section 4.8.3). The interior of the b-barrel harbors the binding site for a (usually single) molecule of ligand, which can be very different in shape, size, and physicochemical properties depending on the structure of the loop scaffold and the cavity of the individual lipocalin. Ligands can be retinoids, bilins, heme, carotenoids, and odorants (not in insects), depending on the physiological context of the carrier protein. However, the physiological ligands are not known from all lipocalins; some may also function without any bound ligand (see Section 4.8.5). The collective name ‘‘lipocalin,’’ proposed in 1987 (Pervais and Brew, 1987), denotes these proteins as carriers of lipophilic ligands that are harbored within a ‘‘calyx’’ (from Greek, meaning cup). Though lipocalins may be found as monomers in vivo and in vitro, they have a tendency to form homomeric oligomers, as frequently observed in the crystalline state. Lipocalins may also complex to other soluble proteins (e.g., retinol-binding protein combines with transthyretrin in vertebrate blood) or to specific cell surface receptors. The latter aspect is not yet well documented, however. There are several other groups of proteins that share a repeated þ1 topology b-barrel with the lipocalins but display significant differences in topology and the construction of the barrel. The lipocalinrelated proteins are represented by the superfamily of fatty acid-binding proteins (see Section 4.8.6),

which were structurally discovered nearly one decade later, and triabin, an unusual single lipocalin that was isolated from an insect even more recently (see Section 4.8.4.3). The specific modifications of the b-barrel structure, as described above for the lipocalins, will be described in the appropriate sections. Two other protein families, the biotin-binding avidins and the bacterial metalloproteinases, which are structurally related to the lipocalins, fatty acidbinding proteins, and triabin, are outside the scope of this chapter dealing with insect proteins. All these variants of b-barrel proteins are part of a structural superfamily, called ‘‘calycins,’’ as they are all characterized by a cuplike cavity as a real or potential internal binding site for a ligand (reviews: Flower et al., 1993, 2000; LaLonde et al., 1994).

4.8.3. Biliproteins: Prototypic Lipocalins 4.8.3.1. Biliproteins with Known Crystal Structure

The green coloration of many insect larvae, thought to provide camouflage at their feeding sites on plants, has been subject to many biological and chemical studies in the past. This green color is known to result from a mixture of yellow carotenoids derived from the food and blue bile pigments synthesized by the insect. Both pigments are usually associated with different proteins, the chromatographic separation of which demonstrates nicely the two underlying coloring principles (review: Kayser, 1985). Insect larvae that are fed artificial diets, usually very poor in carotenoids,

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271

Figure 2 Last instar larvae of Manduca sexta fed tobacco leaves (green form) and artificial diet (blue form), respectively. The difference in color results from the lack of yellow carotenoids in the artificial diet. The blue coloration is due to the presence of insecticyanin, a biliprotein synthesized by the larvae.

consequently exhibit not a green but a blue coloration. Insect scientists working with a laboratory strain of the tobacco hornworm, Manduca sexta, for example, may have rarely seen the ‘‘wild’’ green coloration of the insect in nature (Figure 2). Blue chromoproteins, all most likely representing biliproteins, are present in many insect orders (Kayser, 1985). The eye-catching colors of biliproteins have certainly contributed to the fact they were studied early at the biochemical level and thus had a good chance to contribute to the discovery of the lipocalins. As mentioned above, the third and fourth lipocalin, the structures of which were independently solved by X-ray crystallography, were not only two proteins from insects but also two rather similar ones: the biliproteins from the butterfly Pieris brassicae and from the moth M. sexta (see below). Long before, both were known as proteins with an attractive sky-blue color, which is due to the presence of a bilatrien bile pigment, biliverdin IXg. Both proteins were studied for their crystal structures in the expectation of finding a predominantly a-helical folding like that established for the light-harvesting cyanobacterial C-phycocyanins that represent biliproteins with a covalently bound derivative of biliverdin IXa (Schirmer et al., 1987). 4.8.3.1.1. Bilin-binding protein from Pieris brassicae The biliprotein from P. brassicae, which was given the specific name bilin-binding protein (BBP) (Huber et al., 1987a), was isolated from whole butterflies, where it is located predominantly in the wings, and characterized in the author’s laboratory as a monomeric, nonglycosylated protein

that is noncovalently associated with biliverdin IXg at a 1 : 1 stoichiometry (Zipfel, 1982; Kayser, unpublished data). A molecular mass of 20 kDa was obtained for purified BBP by both gel filtration and sodium dodecylsulfate (SDS)-gel electrophoresis consistent with its monomeric status. Alignment of the BBP sequence revealed closest homology to the biliprotein from M. sexta, known as insecticyanin (see Section 4.8.3.1.2), and to human apolipoprotein D with identities of 43% and 31%, respectively. The mature protein, as isolated from the butterflies, comprises 174 amino acid residues corresponding to a molecular mass of 19 790 Da in agreement with the chromatographic results (Suter et al., 1988). An N-terminal signal sequence of 15 amino acids is found in the precursor, which has been cloned from larvae (Schmidt and Skerra, 1994). In the crystal state BBP was found as a tetramer consisting of two dimers (Huber et al., 1987a, 1987b). The folding of the monomers was that of the typical eight-stranded b-barrel with orthogonal arrangement in the two sheets (Figure 3), as was first described for the retinol-binding protein. A typical short 310-helix is located in the N-terminal sequence just before the first b-strand. A long a-helix is attached along one side of the barrel in the C-terminal region that terminates in a short helical sequence. BBP contains two cysteine bridges that are believed to support this folding structure. Though the sequence homology of these two lipocalins is only 10%, their tertiary structures match almost perfectly in the central and lower segments, while the loop region around the open end of the barrel is more varied. The calyx-like cavity of BBP harbors

272 Lipocalins and Structurally Related Ligand-Binding Proteins

Figure 3 Crystal structure of the bilin-binding protein (BBP) from Pieris brassicae. Upper and lower panels: ribbon drawing with eight b-strands (blue arrows, labeled A–H) and three helices (red, labeled h1–h3). Middle panel: Ca-backbone structure (light blue) of the protein (same orientation as in the upper panel) with bound biliverdin-IXg represented as stick-and-ball model. Oxygens are in red and nitrogens in blue. The N- and C-termini are marked with white dots.

the chromophore ligand, which is not deeply buried in the cavity but located close to the open end of the barrel with the two carboxyl groups of the heme propionate side chains pointing to the solvent. This implies only little contribution of these polar groups to the binding of the tetrapyrrole. The pyrrole nitrogens of the bilin are complexed to water molecules and in contact with the protein via a few hydrogen bonds. The identity of the ligand of BBP was confirmed by its crystal structure as the g-isomer of biliverdin IX. The bilin shows a cyclic helical structure characterized by all-Z configuration and all-syn conformation (Figure 3). This geometry is adopted also by free bilatrienes in solution (Kayser, 1985) and in agreement with the visible absorption spectrum of the bound chromophore (see below). The cyclic helical geometry of a bilatrien gives rise to two enantiomers with opposite helical sense, which are rapidly interconverted in solution. The crystallographic analysis revealed that the protein binds only one enantiomer of the ligand (Figure 3), thus inducing chirality in a compound that is achiral by structure (Huber et al., 1987b; Scheer and Kayser, 1988). The induced optical activity of the P. brassicae BBP is nearly as strong as that in the cyanobacterial phycocyanins, which has a different basis, however. The circular dichroism (CD) spectrum of BBP shows a positive Cotton effect in the visible range, which could be correlated with the righthanded helix of the bilin for the first time (Huber et al., 1987b). Treatment of the holoprotein with urea completely abolishes the optical activity of the holoprotein due to the unfolding of the protein that destroys the specific ligand binding site (Scheer and Kayser, 1988). As shown with the recombinant BBP apoprotein derived from larval cDNA, the holoprotein can be reconstituted by incubation with free biliverdin IXg to yield an apparently native product as judged from its characteristic absorption spectrum (Schmidt and Skerra, 1994). However, no CD spectrum was recorded, hence the correct binding of the chromophore as the right-handed helical enantiomer has not been demonstrated. As indicated above, the visible spectrum of the bound chromophore is similar to that of the free form, as both are in a cyclic helical conformation. Characteristic differences, however, are seen at the long-wave absorption peak with a bathochromic shift of 25 nm and a significant hyperchromic effect to produce an absorbance plateau around 670 nm; the sharp short-wave peak (the typical Soret band of tetrapyrroles) is at 383 nm (Zipfel, 1982; Scheer and Kayser, 1988). The tight fitting of ligand binding to its carrier lipocalin, as is obvious in case of BBP, may

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generally result in some electronic interaction between the bound chromophore and the protein. Conversely, a biliprotein with a fairly unchanged absorption spectrum versus the free bilin is unlikely to represent a lipocalin. 4.8.3.1.2. Insecticyanin from Manduca sexta Insecticyanin from larvae of the tobacco hornworm, M. sexta, was the first insect biliprotein that was purified and studied at the biochemical level (Cherbas, 1973). It can be easily obtained from larval hemolymph, where it may account for up to 5% of the total protein. Insecticyanin is very similar to BBP from P. brassicae with respect to its molecular size (189 amino acids for the protein as isolated; molecular mass of 21 378 Da), the presence of two disulfide bridges, the absorbance spectrum, the noncovalent binding of one molecule of biliverdin IXg per monomer as well as the absence of bound carbohydrate and lipid (Riley et al., 1984; Goodman et al., 1985). The crystal structure of insecticyanin, solved independently and published only weeks after that of BBP, revealed very close similarity to that of retinol-binding protein (Holden et al., 1987). This finding extends the overall similarity between the biliproteins from M. sexta and P. brassicae to the level of their crystal structures, though the sequence identity is only 40%. Moreover, both biliproteins crystallized as tetramers. As shown in Figure 4, insecticyanin is folded as a barrel made up of eight antiparallel b-strands with an orthogonal arrangement of the two sheets, comparable to BBP. The N- and C-terminal sequences with several conserved helical structures are wrapped near the opening of the barrel. There is an extra helical structure inserted between the fourth and fifth b-strand (labeled h2 in Figure 4). The biliverdin ligand is located near the opening with the two propionate side chains directed to the solvent. The ligand of insecticyanin is in contact with hydrophobic residues on both sides, one of which is more polar than the other one in contrast to the situation in the related BBP. Like in the latter, biliverdin IXg in insecticyanin adopts a porphyrinlike cyclic helical conformation, which is also righthanded like that in BBP (cf. Figures 3 and 4). There is no mention of the helical sense in the publication by Holden et al. (1987). As expected from the study of the P. brassicae protein, the chromophore of Figure 4 Crystal structure of insecticyanin from Manduca sexta. Upper panel: ribbon drawing with eight b-strands (blue arrows, labeled A–H) and several helices (red, labeled h1–h5). Middle and lower panels: Ca-backbone structure (light blue) of the protein with bound biliverdin-IXg in the lower panel. The

ligand is represented as stick-and-ball model with oxygens in red and nitrogens in blue. The upper and middle panels show the protein in the same orientation. The N- and C-termini are marked with white dots.

274 Lipocalins and Structurally Related Ligand-Binding Proteins

insecticyanin is optically active, producing a CD spectrum that is consistent with a right-handed helical sense (Kayser, unpublished data).

M. sexta have been studied most thoroughly including their crystal structures, as described above. The possible classification of the other biliproteins as lipocalins depends on the results of future crystallographic analysis or successful modeling of amino acid sequences into the crystal structures of one of the known biliproteins. The full-length amino acid sequences are also known for the two biliproteins from the silk moth Samia cynthia ricini and from two other proteins from the butterfly P. rapae. The latter two proteins, which differ from each other by

4.8.3.2. Biliproteins with Unknown Crystal Structure

Due to their widespread occurrence in insects and easy visibility, a number of biliproteins have been isolated, mostly from lepidopteran species, and biochemically characterized in more or less detail (Table 2). The biliproteins from P. brassicae and

Table 2 Purified insect biliproteins

Insects

Subunit size (kDa)

Oligomeric state

Bilin type (bilin/subunit)

Glyco/ Lipo protein

Reference

Bilin-binding protein BBPb,c Bilin-binding proteins BBP-1, BBP-2 Insecticyaninb,c

19.8

Monomer

BV IXg (1 : 1)

/

Suter et al. (1998)

18

?

BV IXg ? (?)

?/?

Yun (personal communication)

21.4

Tetramer

BV IXg (1 : 1)

/

Riley et al. (1984)

A. convolvuli

21.2

Trimer (?)

BV IXg? (1 : 1)

?/?

22.4

Dimer

BV IXg? (1 : 1)

?/?

Saito and Shimoda (1997) Saito (1997)

I: 20.5 II: 22.7

Monomer Dimer

BV IXg (1 : 1) BV IXg (1 : 1)

?/? ?/?

Saito (1998a)

21.5

Monomer

Phorcabilin (?)

?/?

Saito et al. (1998)

I: Monomer II: Dimer Tetramer ?

I: phorcbilin (?) II: BV IXg (?) unknown (?)

?/? ?/? ?/?

Saito (1998b)

Name of protein a

Lepidoptera Pieris brassicae Pieris rapae

Manduca sexta Agrius convolvuli Attacus atlas

insecticyanind Biliverdin-binding protein (BBP)d Biliverdin-binding proteins BBP-I,c BBP-IIc Biliprotein BP ¼ P-IId Biliproteins BP-I,d BP-IId

Samia cynthia ricini Antheraea yamamai Rhodinia fugax Cerura vinula

Cerura (vinula)

I: 22.6 II: 22.9 80

Heliothis zea

biliprotein Blue chromoprotein

150

Tetramer

BV (?)

þ/þ

Trichoplusia ni

Chromoprotein

150

Dimer

BV (?)

þ/þ

Spodoptera litura

Biliverdin-binding proteins BP-1–BP-4 Biliverdin-binding vitellogenin

165

Dimers

BV (?)

þ/þ

188

Dimer

BV (?)

?/þ

Maruta et al. (2002)

Cyanoprotein

83

Tetramer

BV (2 : 1)

þ/

Chino et al. (1983)

Blue lipophorin

250/80/20

>700 kDa

BV IXg (?)

?/þ

Cyanoproteins CP1–CP4

76

Hexamers

BV IXg (4 : 1)

þ/

Haunerland et al. (1992b) Chinzei et al. (1990)

Spodoptera litura

Scheer and Kayser (1988) Haunerland and Bowers (1986) G. Jones et al. (1988) Yoshiga and Tojo (1995)

Orthoptera Locusta migratoria

Heteroptera Podisus maculiventris Riptortus clavatus a

As referred to in the reference. Crystal structure available; see Table 1. c Full sequence available; see Table 1. d N-terminal sequence available. BV, biliverdin. b

Lipocalins and Structurally Related Ligand-Binding Proteins

a few residues only, are of the same size as BBP from P. brassicae to which they show 94% identity (Yun, personal communication). Hence, it is most likely that the biliproteins from P. rapae have crystal structures almost identical to that of BBP from the closely related P. brassicae. N-terminal sequences are known from several moth biliproteins (see Table 2). The chromophores of most insect biliproteins have been described on the basis of their visible absorption spectra as either biliverdin IXg or ‘‘biliverdin(-like)’’ without further specification (Table 2). A more rigorous approach by, for example, microchemical degradation has not been performed with any of these chromoproteins. On the other hand, the identification of biliverdin IXg, though tentative so far, in most of the lepidopteran proteins is not surprising since this biliverdin isomer is typical for this insect order (review: Kayser, 1985). Phorcabilin, a derivative of biliverdin IXg with extended chromophore geometry, seems to be associated with proteins in two saturniid moths only, in accordance with the restrictive occurrence of this bilin (Kayser, 1985). The ligand of the biliprotein from the moth Cerura vinula has not yet been identified. Preliminary results suggest a new chromophore, possibly a derivative of biliverdin IXg (Scheer and Kayser, 1988; Kayser, unpublished data). In many cases, the names of the insect biliproteins that have been described to date may give rise to confusion, because they are used for different proteins, or they simply refer to the blue color, as documented in Table 2. Though this is no attempt to propose a nomenclature for biliproteins, it should be kept in mind that the name ‘‘bilin-binding protein’’ or ‘‘BBP’’ has been given specifically to the biliprotein(s) from P. brassicae (Huber et al., 1987a, 1987b). Similarly, ‘‘insecticyanin’’ has been coined for the biliprotein from M. sexta (Cherbas, 1973). These names should not be used for biliproteins from other species. Furthermore, a protein may be denoted as ‘‘biliverdin-binding’’ only if the chromophore has been definitely identified as one of the isomers of biliverdin IX. If the chemical class of the (blue) ligand is unknown, the protein may be described just as, for example, a ‘‘blue chromoprotein,’’ ‘‘blue lipophorin,’’ or ‘‘cyanoprotein,’’ as in the past. To avoid confusion in future work it is proposed to use the generic term ‘‘biliprotein,’’ abbreviated as ‘‘BP,’’ linked to the name of the species from which it was isolated. At least two groups of biliproteins may be discriminated on the basis of their overall characteristics (Table 2). One group is characterized by small proteins of 20 kDa, lack of sugar and lipid

275

constituents, binding of a single ligand per subunit, and occurrence as monomers or as homo-oligomers. This group may represent lipocalins, as typified by BBP from P. brassicae and insecticyanin from M. sexta. The second group comprises proteins characterized by much larger subunits in the 80 kDa or even 150 kDa range, the presence of sugar and/or lipid and the aggregation to complexes with sizes of up to 600 kDa and more. Some proteins of this type, such as the biliverdin-binding vitellogenin from Spodoptera litura (Maruta et al., 2002), the blue lipophorin from Podisus maculiventris (Haunerland et al., 1992b), and the cyanoprotein from Riptortus clavatus (Chinzei et al., 1990) show properties that relate them to the storage proteins which are used during the embryo and metamorphosis stages (see Haunerland, 1996). In most of the biliproteins listed in Table 2, the absorption spectrum of the bound chromophore differs more or less significantly from that of the free ligand obtained after extraction with an organic solvent. As in the Pieris BBP, the long-wave absorption maximum of the bilin is usually redshifted (bathochromic effect) and increased (hyperchromic effect), which implies some electronic interaction between the ligand and the protein. Whether such spectral changes indicate that the ligand is specifically bound with respect to its structure and the binding site is not clear. It may well be that less specific binding of a ligand to a site on the protein surface or to the lipid moiety of a protein also results in a shift in the absorption spectrum. In any case, the binding cavity of the known lipocalins would not be large enough to harbor more than one bilin per monomer. So, the nature of the ligand-binding site of the large-size biliprotein remains conjecture. However, the crystal structure of the 80 kDa subunit biliprotein from Cerura vinula, isolated in the author’s laboratory, may be solved in the near future if current attempts to obtain high-quality crystals are successful. 4.8.3.3. Molecular Variants of Biliproteins

In a number of species, multiple biliproteins have been isolated and characterized by their amino acid composition, N-terminal sequence, and molecular mass (Table 2). Examples are found among the small-size (20 kDa) proteins from two saturniid moths. In Samia cynthia ricini, the two biliverdinbinding proteins (BBP-I and BBP-II) from larval hemolymph share only 48% identity in their N-terminal 50 amino acid residues. BBP-II is likely identical to the biliprotein isolated from the molting fluid at pupation from the same insect (Saito, 1993, 1998a). The identity of the predominant BBP-I from this moth with BBP from P. brassicae

276 Lipocalins and Structurally Related Ligand-Binding Proteins

and insecticyanin from M. sexta is 46% and 52%, respectively. The corresponding value for BBP-II is 42% for both comparisons. In Rhodinia fugax, the biliproteins BP-I from larval hemolymph and cuticle and BP-II from epidermis are similar to one another and to those from P. brassicae and M. sexta, respectively (Saito, 1998b). An example for multiple large-size (165 kDa) proteins has been found in Spodoptera litura, from which four dimeric hemolymph biliverdin-binding proteins, differing in their isoelectric point, have been isolated indicating the presence of several nonidentical subunits of about equal size (Yoshiga and Tojo, 1995). Unfortunately, partial sequences are not available to support this conclusion, but immunological differences have been recognized. As mentioned above, another biliverdin-binding protein from the same moth has been identified as a vitellogenin as it is female-specific and shows significant sequence homology with lepidopteran vitellogenins (Maruta et al., 2002). The oligomeric situation with the cyanoproteins from the bug Riptortus clavatus is different (Chinzei et al., 1990). Here, two distinct subunits of about equal size combine to form four biliverdin-associated hexameric complexes which have been isolated as CP1 to CP4. According to their cDNAdeduced amino acid sequences, these proteins are closely related to arthropod hemocyanins and phenol oxidases, which are members of the superfamily of hexamerins (Miura et al., 1998). Their hexameric state and the high ratio of chromophore per subunit of 4 : 1, as calculated for CP1, support the conclusion that these bug cyanoproteins are not lipocalins. Full data at the gene level on molecular variants of biliproteins have been obtained for the established lipocalin from M. sexta (Kiely and Riddiford, 1985; Riddiford et al., 1990; Li and Riddiford, 1992, 1994, 1996). In this insect, two major isoelectric forms of insecticyanin, described as a more acidic INS-a (pI 5.5) and a more basic INS-b (pI 5.7), have been identified. Both forms are present in the larval epidermis and cuticle, whereas only INS-b is found in the hemolymph. It is this form which has been studied as the first insect biliprotein by Cherbas (1973). The protein sequences encoded by the cDNAs for INS-a and INS-b, respectively, differ in 13 of the 189 amino acid residues of the mature protein; the N-terminal signal sequences are identical. The 30 noncoding regions of the two cDNAs contain a sequence stretch that is unique for each gene and led to specific probes for expression studies of the two duplicated insecticyanin genes. Comparable to the results on insecticyanin in M. sexta, two isoelectric forms of the BBP were

purified from P. brassicae with pI values of 6.4 (BBP-I) and 6.2 (BBP-II), respectively (Kayser, unpublished data). The two proteins differ only in the N-terminal residue, which is asparagine in the predominant form BBP-I and aspartate in the minor variant BBP-II (Huber et al., 1987b). This single difference in sequence fully accounts for the observed difference in the isoelectric point of the two forms. As expected, the crystal structures of the two BBP variants are identical, but BBP-II does not dimerize. In contrast to the two isoforms obtained as proteins from adult insects, only one cDNA sequence coding for BBP (174 amino acid residues of the mature protein plus an N-terminal signal peptide of 15 amino acids) has been found in last instar larvae, which are actively expressing this gene (Schmidt and Skerra, 1994), and accumulate most of the BBP holoprotein during the insects’ life (Kayser, 1984; Kayser and Krull-Savage, 1984). Hence, BBP-II may arise artifactually by deamidation of BBP-I during the process of purification. The presence of the minor form BBP-II in vivo has yet to be confirmed. Two isoelectric forms of biliproteins are also known from the related butterfly, P. rapae, which differ in 12 nucleotides in their cDNA sequences (Yun, personal communication). Hence, there are obviously separate genes for the two biliproteins in P. rapae, in contrast to only a single gene in P. brassicae. 4.8.3.4. Developmental Expression and Biosynthesis of Biliproteins

Though biliproteins are widespread among insects and may share a number of physicochemical features, there seems to be no clear common pattern with respect to their location in the body, their site of synthesis, and their fate during development. Detailed studies of the transcription of genes coding for apobiliproteins and the corresponding synthesis of proteins have been performed for insecticyanin in M. sexta (Riddiford, 1982; Trost and Goodman, 1986; Riddiford et al., 1990; Li and Riddiford, 1994). Insecticyanin is present in the larval epidermis and hemolymph up to the adult stage. The highest concentration is found in the hemolymph of early fourth instar larvae and in adults after eclosion (up to 0.8 mg ml1), while the levels decrease after each larval molt. The genes ins-a and ins-b coding for the two isoelectric forms of insecticyanin are expressed during all larval instars with significant cyclic variations, followed by corresponding increases and decreases of the protein titers. Since the expression of the ins genes is restricted to the larva, insecticyanin present in the adult insect must

Lipocalins and Structurally Related Ligand-Binding Proteins

p0130

be retained from the larval stage. Both insecticyanin variants are mainly synthesized in the larval epidermis and to a lesser extent in the fat body. While the epidermis stores both proteins in granular form and secretes them into the cuticle, it delivers only INS-b to the hemolymph. INS-b is also produced as the predominant isoform in the fat body, but obviously released quickly and not stored there, as this tissue lacks blue coloration. The epidermal expression of ins-a lasts until pupal commitment, while that of ins-b continues up to the wandering stage. This differential control of gene expression is due to a small peak in the ecdysteroid titer occurring after the disappearance of juvenile hormone (Riddiford et al., 1990; Li and Riddiford, 1994). Thus, the developmental regulation of insecticyanin synthesis is apparently part of the overall endocrine program that controls the larval–pupal transition. In the black mutant of M. sexta, which is characterized by a lack of juvenile hormone (see Chapter 3.7), the expression of both ins genes is reduced in the epidermis, while it is enhanced in the fat body, suggesting a direct, tissue-specific effect of juvenile hormone on the expression of these genes (Li and Riddiford, 1996). Insecticyanin is present in mature eggs of M. sexta at a concentration exceeding that in the hemolymph, though the ins genes are not expressed in any cell type in pupae and adults. The protein persisting from the larval stage is taken up from the hemolymph into the developing oocytes by a saturable and specific membrane-bound mechanism that requires the presence of calcium, as found by Kang et al. (1995, 1997). An apparently multimeric receptor with an estimated size of 185 kDa that selectively binds insecticyanin with high affinity (Kd 17 nM) has been characterized in detergent extracts from oocyte membranes. Thus, insecticyanin in the Manduca egg behaves like the storage proteins vitellogenin and lipophorin despite the differences in protein architecture, molecular size and mechanism of uptake into the egg (review: Haunerland, 1996) (see Chapter 3.9). The expression of the BBP in P. brassicae is under developmental and tissue-specific control. Its mRNA can be demonstrated in penultimate and last instar larvae with a depression during the last larval and larval–pupal molts (Schmidt and Skerra, 1994). This kind of cyclic expression resembles that seen for insecticyanin in M. sexta. On the other hand, the BBP gene is not expressed in the larval epidermis but in the fat body, predominantly in the dorsolateral sheets and in the fat body tissue underlying the epidermis. This is not surprising since this tissue is of bluish coloration. Only little

277

BBP mRNA is seen in the ventrolateral fat body, which is colorless indicating that the holoprotein is not stored there to a significant extent. Furthermore, the gene seems to be active also in the larval ‘‘gonads’’ (testes, according to this author). In marked contrast to the situation in M. sexta, the expression of the BBP gene in P. brassicae is again strong in the late pupa (pharate adult) and in young adults before it ceases almost completely (Schmidt and Skerra, 1994). This developmental pattern of BBP expression is in accordance with results from biochemical studies on the protein titer and the incorporation of specific precursors into the bilin and the apoprotein, respectively (Kayser, 1984; Kayser and Krull-Savage, 1984). The most active phases of holoprotein synthesis are in the last larval instar before the wandering stage and in late pupae following the increased ecdysteroid titer, which triggers adult development (see Chapter 3.5). The BBP synthesis ceases 2 days after adult emergence. The strong synthesis of BBP in developing adults is surprising as it is not sequestered into the eggs of P. brassicae, in contrast to the fate of insecticyanin in M. sexta. Most of the BBP in the butterfly is located in the wings, which are blue underneath the layer of white scales containing large amounts of pterin pigments in granular form (review: Kayser, 1985). Recent studies in the author’s laboratory revealed that this wing BBP is synthesized de novo as holoprotein in the wings during adult development, while the BBP gene is switched off in the rest of the body (Sehringer, 1999; Sehringer and Kayser, unpublished data). These studies also demonstrated that the wings are capable of synthesizing the bilin ligand of BBP from the specific heme precursor 5-aminolevulinate (see below). Results from a developmental study of the activity of one of the key enzymes of tetrapyrrole synthesis, porphobilinogen synthase, are in accordance with the prominent phases of BBP synthesis and with the wings as the sites of BBP synthesis in the pupal stage (Kayser and Rilk-van Gessel, unpublished data). There is an interesting difference between P. brassicae and P. rapae relating to the regulation of their biliproteins during development. In P. rapae, the isoform BBP-1 is abundant in the larval hemolymph, while there is more BBP-2 in the pupal stage (Yun, personal communication). The site of synthesis of the two P. rapae biliproteins remains to be studied. No isoforms with differential temporal expression are known from the closely related P. brassicae, as mentioned above. In the larvae of Spodoptera litura, the four biliverdin-binding proteins that are composed of two 165 kDa lipoprotein subunits (Table 2) also

278 Lipocalins and Structurally Related Ligand-Binding Proteins

show hemolymph levels that fluctuate cyclically in the course of larval development (Yoshiga et al., 1998), as described above for insecticyanin. The timing of BP-4 differs from that of the others, as it appears in earlier instars and has a low titer in the last instar, when the levels of the others are high. Synthesis of BP-4 seems to be triggered by juvenile hormone (Yoshiga and Tojo, 2001). At pupation, the biliproteins are taken up by the fat body, which turns blue in the pupa. A comparable sequestering of large-size biliproteins as glycolipoproteins by the fat body from the hemolymph at pupation is observed in the moth Heliothis zea (Haunerland and Bowers, 1986) and in the bug Riptortus clavatus (Chinzei et al., 1990). These biliproteins with subunits of 150 kDa and 76 kDa, respectively, represent high-density lipoproteins (see Chapter 4.6) and behave like storage proteins that are used up for adult development and egg formation. It is remarkable that the Manduca insecticyanin, a typical 20 kDa lipocalin, also behaves like a storage protein in the embryo, as described above. This demonstrates that the presence of a bilin ligand does not denote a specific functional class of proteins. As for the biosynthesis of the bilin chromophore of biliproteins, labeling studies in a number of insects have demonstrated that the heme precursor 5-aminolevulinate is efficiently incorporated into the bilin (review: Kayser, 1985). More detailed experiments, also in relation to the synthesis of the apoprotein, seem to be available only for the BBP from P. brassicae (Kayser, 1984; Kayser and Krull-Savage, 1984). These studies revealed that the syntheses of the bilin and the apoprotein are coordinated but independent from each other, since cycloheximide has a strong inhibitory effect on the synthesis of the apoprotein, while the incorporation of radiolabel from 5-aminolevulinate into protein-bound bilin was only weakly reduced. This suggests the presence of a relatively large pool of apoprotein available for binding of the heme derivative. A study of the time course of BBP synthesis in young adults of P. brassicae demonstrated further that label from [14C]5-aminolevulinate appeared in the bilin almost immediately with no lag phase, comparable to the results for the apoprotein labeled with [3H]leucine (Kayser and Krull-Savage, 1984). This result is consistent with a straightforward synthesis of the bilin, presumably via a heme precursor. Obviously, the bilin is not accumulated like a degradation product of some relatively stabile heme protein, for example like hemoglobin in vertebrates. The formation of bilins from heme has not yet been studied at the enzyme level in insects, though the action mechanism of heme oxygenase is fairly

well understood and evolutionarily conserved (reviews: Maines, 1997; Wilks, 2002). It is furthermore unknown whether apobiliproteins or their precursors could serve as heme oxygenases, or whether the open chain tetrapyrrole is synthesized independently to associate with the apoprotein, as studies in P. brassicae suggested (Kayser and Krull-Savage, 1984). This possibility gains some support from the apparently successful reconstitution of BBP from its recombinant apoprotein (Schmidt and Skerra, 1994). 4.8.3.5. Putative Functions of Biliproteins

Usually, biliproteins are considered to contribute primarily to camouflage coloration of insects, in particular to reduce the risk for larvae feeding on (green) plants of being preyed upon. This is due to their strong blue color, which combines with the yellow color of carotenoids derived from the food plant to yield green coloration that may be modified by, e.g., dark pigmentation, to further enhance any camouflage effect (review: Kayser, 1985). This role is certainly true and evident in many insects, as in the example shown in Figure 5. However, there are also examples where biliproteins are present in epidermis, hemolymph, and/or fat body of the larva but are masked by other pigmentation of the cuticle. One obvious example is the butterfly P. brassicae. The larval integument displays a pattern of black and yellow pigments that mask the biliprotein underneath, and in the adults, where the bulk of

Figure 5 Fourth instar larva of Cerura vinula as an example for camouflage coloration based on yellow carotenoids and a blue biliprotein present in epidermis and hemolymph, combined with ‘‘lytic’’ dark pigmentation in the cuticle.

Lipocalins and Structurally Related Ligand-Binding Proteins

the biliprotein is located in the wings, the scales are bright white, except for a few black patches. Presumably, the camouflage potential is a secondary property of biliproteins that is successfully utilized in many insects. A major, more general role of biliproteins is inferred from several considerations. The high cost of establishing the complex multienzyme pathway of heme synthesis followed by heme cleavage seems to be inadequate for biliproteins to be limited to just a role in camouflage, which might be achieved by other, ‘‘cheaper’’ means. Furthermore, the strong synthesis of heme products without any known need for a corresponding large quantity of a heme protein, such as hemoglobin or cytochrome P450 (see Chapter 4.1), suggests a genuine and vital role for the biliproteins as end (not waste) products of the heme pathway. In newly emerged adults of P. brassicae, for example, about 90% of the heme precursor 5-aminolevulinate is allocated directly to the synthesis of BBP, even at the time of peak formation of mitochondrial cytochromes during flight muscle development (Kayser, 1984). The photochemical properties of the cyanobacterial biliproteins, which function as accessory light-harvesting complexes, led to a study of the photochemical behavior of an insect representative, BBP from P. brassicae (Scheer and Kayser, 1988; Schneider et al., 1988). The fluorescence intensity of BBP was very weak, compared to that of the cyanobacterial counterparts, and there was no indication of an intermolecular energy transfer in accordance with the monomeric nature of this protein. Overall, the photochemical properties of BBP are much too weak to support any speculation about a possible role based on the use of light for energy conversion or signaling. Most likely, this is true for insect (animal) biliproteins in general. Concerning possible metabolic roles of biliproteins, it must be recalled that the cleavage of heme yields an open-chain tetrapyrrole, which is a biliverdin in all studied organisms, as well as carbon monoxide and iron in equimolar quantities. All these products are biologically active. A role for the released iron is difficult to reconcile since it has to be introduced into the porphyrin for its cleavage via its Fe-complex, heme. The production of carbon monoxide is possibly more relevant, as evidence is being accumulated for this gas to function as a second messenger comparable to the well-established role of nitric oxide which regulates a wide spectrum of biochemical and physiological processes by binding to the heme moiety of soluble guanylyl cyclases. Moreover, the pathways of nitric oxide and carbon monoxide signaling have been shown

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to interact (reviews: Baran˜ ano and Snyder, 2001; Hartsfield, 2002; Ryter et al., 2002). Thus, heme oxygenase is in a position to potentially control a number of cellular reactions. Isoforms of heme oxygenases with different functions and tissue specificity are known from vertebrates. Insects have been widely neglected in this field so far. There are more speculative roles for bilins and their carrier proteins. The cleavage of the heme ring to an open-chain bilin is unlikely to be just a means to control the cellular levels of porphyrins because of the high investment to establish such a mechanism. Multistep biosynthetic pathways are usually regulated at the level of initial (key) enzymes, not at the terminal ones. Porphyrins are cytotoxic due to their photochemical properties and therefore have to be inactivated or eliminated. Their cleavage to a linear tetrapyrrole requires the insertion of ferrous iron for the activation of molecular oxygen and reduction equivalents, usually NADPH. However, bilins may also have regulatory roles according to recent studies in vertebrates. For example, biliverdin has been shown to inhibit soluble guanylyl cyclase in vivo and in vitro (Koglin and Behrends, 2002), to play a role in embryonic development (Falchuk et al., 2002), and, as a redox partner of bilirubin, to play a vital role in the prevention of cellular damage by reactive oxygen and nitrogen species (Baran˜ ano et al., 2002; Kaur et al., 2003). Apparently, a variety of actions of linear tetrapyrroles in biological systems are just emerging.

4.8.4. Nitrophorins and Related Proteins 4.8.4.1. The Problem of Blood Feeders

The basic lipocalin fold has evolved as a universal scaffold that can be modified to achieve affinity for chemically diverse ligands involved in a variety of biological functions. This universality becomes strikingly evident with the nitrophorins and functionally related proteins that have evolved independently in several arthropods specialized in taking blood meals. General aspects of this special feeding behavior have been reviewed very recently (Ribeiro and Francischetti, 2003). Blood-sucking arthropods, such as a number of bugs and ticks, have developed biochemical means to maintain a continuous blood flow at the feeding site and to prevent or delay any counteraction of the host’s body that would reduce feeding success. This common requirement of blood feeders obviously guided the evolution of proteins specialized to interact at different sites and in different ways with the complex

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process of blood coagulation in vertebrates. Obviously, the lipocalin fold provided a versatile basis to create binding sites for powerful ligands acting together to support the insects’ needs. The basic studies on this group of functionally highly specialized lipocalins have been performed in Rhodnius prolixus, a South American bug known to transmit Trypanosoma cruzi, the vector of Chagas’ disease. A general overview with a focus on the structures of the specialized proteins of blood-feeding insects has been given by Montfort et al. (2000). The chemical and physical properties of nitrophorins have been reviewed by Walker et al. (1999). 4.8.4.2. Crystal Structures of Nitrophorins from Rhodnius prolixus

The nitrophorins of the bug R. prolixus are stored in the salivary gland, which therefore looks red, and are injected, together with a number of other proteins of related functions, into the blood of the host. There are four nitrophorins, NP1 to NP4, that have been purified from the insect tissue, cloned, and produced as recombinant proteins for crystallization studies (Champagne et al., 1995; Ribeiro et al., 1995; Andersen et al., 1997; Sun et al., 1998). The numbering of the nitrophorins is in the order of their relative abundance with NP1 representing the predominant one. Each protein has a mass of 20 kDa as is typical for members of the lipocalin family. In pairwise comparisons, NP1 and NP4 show 90% sequence identity, while this is 80% for NP2 and NP3. The identities between the two pairs are much lower, about 45%. This demonstrates different degrees of relatedness suggesting that the four nitrophorins originated from two gene duplications. The overall sequence identity of the nitrophorins is 38%. The crystal structures of the nitrophorins, which are available from NP1, NP4, and NP2 in various complexes, are largely identical (Andersen et al., 1998; Weichsel et al., 1998, 2000). All of them show the typical conserved lipocalin folding of a b-barrel consisting of eight antiparallel b-strands with orthogonal orientation in the two sheets (Figure 6), as described in detail for the biliproteins from P. brassicae and M. sexta (see Section 4.8.3.1). Moreover, the nitrophorins share with the biliproteins the conserved long helix (actually two helical regions here, labeled h2 and h3 in Figure 6) at the extended C-terminal sequence that does not contribute to the barrel structure. Furthermore, two disulfide bridges are present at positions comparable to those in the biliproteins. The main difference between the two types of lipocalins refers to the ligand, which is heme in the ferric state in each of the nitrophorins (Figure 7). The heme is held in place

Figure 6 Crystal structure of nitrophorin 1 from Rhodnius prolixus. Ribbon drawings with eight b-strands (blue arrows) and several helices (red, labeled h1–h3). The N- and C-termini are marked with white dots. For the heme ligand, see Figure 7.

via 10 hydrophobic amino acid side chains and a histidine (His59) as the fifth iron ligand. This detail is reminiscent of the hemoglobins in which the heme is in the ferrous state, however. Even more, these two types of hemoproteins represent quite different protein structures: b-barrels in the nitrophorins and a-helical globular folds in the hemoglobins, demonstrating that they are evolutionarily unrelated. In the nitrophorins, the structure of the heme is very unusual as it is distorted and nonplanar. The carboxyl groups of the two propionate side chains of heme are hydrogen-bonded to the same lysine, and one of the propionates is also linked by an unusual carboxylate–carboxylate hydrogen bond to an aspartate side chain. Both heme propionate groups point to the solvent (Figure 7). The orientation of the heme in the nitrophorins is thus comparable to that of the cyclic but open-chain bilins in the biliproteins, as is

Lipocalins and Structurally Related Ligand-Binding Proteins

Figure 7 Crystal structure of nitrophorin 1 from Rhodnius prolixus. Ca-backbone structures (light blue) with the heme ligand in complex with nitric oxide (upper and middle panels) and histamine (lower panel), respectively. Heme is represented as stick-and-ball model with oxygen in red, nitrogen in blue, and iron in green. Two helices (h1 and h2) and the C-terminus are labeled. For further orientation, compare to Figure 6.

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the location of the heme near the opening of the barrel. Though the nitrophorins show a higher degree of sequence identity with the biliproteins, their core lipocalin structures fit better to the vertebrate lipocalins. On the other hand, the positions of the disulfide bridges are identical in the two groups of insect lipocalins, while they are different from the mammalian lipocalins. Evolutionary relatedness in this case may hence be better reflected by the disulfide pattern than by identical positions of the core atoms of the b-barrel. As can be seen from the crystal structure depicted in Figure 7, the nitrophorins are isolated as complexes of the heme iron with nitric oxide, which can be exchanged by histamine (Weichsel et al., 1998). The heme is in the low-spin state in the complexes with nitric oxide and histamine (Soret maximum at 419 nm and 413 nm, respectively), and in the highspin state when these ligands are replaced by water (Soret maximum at 404 nm). The dissociation constants (Kd values) of nitric oxide in complex with the various ferric-heme nitrophorins is in the 10  109 M to 1000  109 M range, depending on pH and the protein isoform. The NP2/NP3 pair shows higher affinities than the NP1/NP4 isoforms. In NP4, binding of nitric oxide changes the conformation of two loops at the b-barrel’s opening so that nitric oxide gets enclosed by hydrophobic residues after several water molecules have been expelled from the cavity (Weichsel et al., 2000). Keeping the heme iron in the oxidized (ferric) state is very important since the nitric oxide complex with ferrous iron is more stable by about six orders of magnitude, which means practically irreversible binding. The ferric state of heme is stabilized by the protein structure. The affinity for the heme ligands is modulated by pH in such a way that at the weakly acidic pH (5) of the salivary gland the nitric oxide–heme complex is more stable than at the higher pH (7.5) at the host’s feeding site. This pH difference facilitates both the release of nitric oxide into the host’s blood and the binding of histamine, due to its higher affinity, in place of nitric oxide. Histamine binds with a Kd of 20  109 M at the host’s pH, which is a 100-fold higher affinity compared to that of nitric oxide in the same environment. NP2, also known as prolixin-S, also exerts anticoagulant activity in addition to its nitric oxide and histamine binding properties. Though details are not known, NP2 interferes at an early step with the multilevel cascade of blood coagulation. Very recently, two additional proteins most likely representing new nitrophorins have been identified in the salivary gland of R. prolixus, and named NP5 and NP6 (Moreira et al., 2003). One of these new

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proteins, NP5, has been partially characterized as a heme protein, according to its absorption spectrum, with an N-terminal sequence revealing high similarity to NP4. Furthermore, NP5 binds nitric oxide and shows a spectral shift of the Soret peak from 404 nm to 422 nm upon binding similar to that of the other nitrophorins. The other protein, NP6, is only tentatively described as a nitrophorin so far. Interestingly, the relative abundance of the nitrophorins in the salivary gland depends on the life cycle stage of this bug (see Section 4.8.4.5). It may be of interest to note that the bedbug Cimex lectularius, a blood feeder like R. prolixus but a member of a different family of Hemiptera, also has a heme-based donor protein for nitric oxide (Valenzuela and Ribeiro, 1998). By this property, it may also be called a nitrophorin. However, its amino acid sequence is unrelated to those of the Rhodnius nitrophorins and shows no lipocalin signature. It will therefore not be discussed further here. These nonhomologous but functionally equivalent proteins have likely evolved independently to meet the common constraints of blood feeding. 4.8.4.3. Other Lipocalins from Blood-Feeding Insects

The anticoagulant activity of the nitrophorin NP2 from R. prolixus has already been mentioned above. Another protein supporting the action of the nitrophorins as donors of nitric oxide to the host has been isolated and cloned from the salivary gland of this bug (Francischetti et al., 2000). This protein, called Rhodnius prolixus aggregation inhibitor 1 (RPAI-1) or salivary platelet aggregation inhibitor 1, comprises 155 amino acids in its mature form and has a molecular mass of 19 kDa. Though its crystal structure is not known, it obviously represents also a lipocalin as suggested by its sequence homology with some other insect proteins believed to have a lipocalin structure. These are pallidipin and triabin (see below) from the bug Triatoma pallidipennis, which both interfere with blood coagulation (Noeske-Jungblut et al., 1994, 1995). RPAI-1 has a nucleotide binding site specific for ADP (and other adenine nucleotides) that is known to potentiate platelet aggregation induced by agonists such as collagen (Francischetti et al., 2002). RPAI-1 is suggested to be most efficient in scavenging small concentrations of ADP that would otherwise induce the formation of large platelet aggregates. The sequence of a related salivary platelet aggregation inhibitor 2 has been reported also from R. prolixus (Champagne et al., 1996). Quite recently, another protein has been detected in the saliva of R. prolixus that apparently supports

the bug in taking a blood meal. The new protein binds biogenic amines that are known to promote platelet aggregation in concert with ADP (Andersen et al., 2003). This biogenic amine-binding protein (ABP) binds norepinephrine with highest affinity (Kd 25 nM), followed by serotonin (Kd 100 nM) and epinephrine (Kd 350 nM). Alignment of the amino acid sequence of ABP with those of the four nitrophorins revealed significant similarity, a little lower than that between the nitrophorins. The lipocalin nature of ABP is further stressed by four cysteines at comparable positions (Figure 8). ABP is not associated with heme, which is explained by the absence of the proximal histidine that acts as fifth ligand of the heme iron in the nitrophorins. Hence, ABP lacks the structural basis for nitric oxide binding. Triabin from Triatoma pallidipennis, a bloodsucking bug like R. prolixus, is a 16 kDa protein consisting of 142 amino acids in the mature form that inhibits thrombin by forming an equimolar noncovalent complex with this key compound for blood coagulation (Noeske-Jungblut et al., 1995). Its structure is remarkable (Figure 9), as it turned out to be an unusual lipocalin with an eight-stranded b-barrel in which, however, the strands B and C of the first sheet are exchanged resulting in an ‘‘up–up–down–down’’ topology instead of the strict antiparallel ‘‘up–down–up–down’’ strand orientation (Fuentes-Prior et al., 1997). Triabin lacks the long C-terminal a-helix that terminates in a short b-strand in typical lipocalins, which accounts for its lower size. Instead short helices are found at both termini. Like the retinol-binding proteins but unlike the biliproteins and nitrophorins, triabin contains three disulfide bridges. A sequence aligment of triabin and nitrophorins, together with related proteins, is shown in Figure 8. 4.8.4.4. Lipocalins from Blood-Feeding Ticks

The common feeding biology of blood-feeding insects and ticks has resulted in the convergent evolution of proteins serving the same goal, which is to support blood feeding by counteracting the hosts’ responses to the attack. These functions are obviously best performed on the basis of a common protein structure, which turned out to be the conserved lipocalin fold. The protein structures as well as their ligands have been studied in two ticks, Rhipicephalus appendiculatus, which prefers cattle, and Dermacentor reticulatus, feeding on rodents. Three histamine-binding proteins (HBP1 to HBP3) were identified in salivary glands of R. appendiculatus and cloned, and the crystal structure of one of them was solved (review: Paesen et al., 2000).

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Figure 8 Aligment of the nitrophorins 1 and 2 (NP1, NP2) and biogenic amine-binding protein (ABP) from Rhodnius prolixus, triabin (TRI) from Triatoma pallidipennis and Bla g 4 (Bg 4) from Blattella germanica. N-terminal secretion signals are boxed in gray. Residues typically conserved in lipocalins are boxed in black. In NP1 and NP2, the heme-binding His59 are also boxed in black. Asterisks indicate identical residues, double and single points more or less conserved substitutions.

Remarkably, these proteins are sex-specific, as two forms, HBP1 and HBP2, are found only in females, while HBP3 is restricted to males. Moreover, while the female-specific proteins are present only in adult ticks and produced only during the early phase of the single feeding period of females, the male-specific form is not restricted to the adult stage but occurs also in nymphs and larvae, and is synthesized over the entire feeding period involving several attacks of a host. These differences suggest that the three forms of HBPs are adapted to the sex-specific feeding biology of this tick species and may serve different needs. According to alignment studies of the amino acid sequences, the two female-specific isoforms, HBP1 and HBP2, are more closely related to each other (66% identity) than to the male protein, HBP3 (32% and 39% identity, respectively). The HBP sequences are unrelated to those of the nitrophorins and other lipocalins. The three structurally conserved regions of the lipocalins are also found in the HBPs, though as substantially modified motifs not readily recognized in the sequences. The molecular masses, based on the sequences without the N-terminal signals, are 19.5 kDa for both HBP1 and HBP2, and 21 kDa for HBP3. There are more sex-specific differences between these proteins: the male protein HBP3 is glycosylated and secreted as a dimer apparently linked by a disulfide bond, while HBP1 and HBP2 from females are monomeric and not posttranscriptionally modified. All HBPs, studied as recombinant proteins, bind histamine with high specificity but different affinities that are not related to sex, however. HBP1 binds histamine with

a Kd of 18  109 M, while the values for HBP2 and HBP3 are in the 1–2  109 M range. The crystal structure of HBP2 has been reported (Paesen et al., 1999). The overall structure matches the lipocalin fold with eight-stranded antiparallel sheets arranged to a fairly spherical b-barrel (Figure 10). This basic fold is modified by several structural details that are unique to this protein. Nevertheless, the structure of HBP2 is most similar to that of the BBP from the butterfly, P. brassicae (see Section 4.8.3.1.1). The N-terminal sequence of HBP2, comprising two helical structures, is extended and attached to the b-barrel via hydrogen bonds. The second helix, labeled h1 in Figure 10, occludes the open end of the barrel. The long C-terminal region comprises another helix, labeled h2, which is fixed to the barrel via two disulfide bridges. The most characteristic feature of HBP2 refers to the barrel cavity, which offers two distinct binding sites for two molecules of histamine (Figure 10), whereas a single ligand is usually present in classical lipocalins. The two sites are located at opposite ends of the cavity, which is separated into two parts due to the side chains of a tyrosine, a glutamate, and a tryptophane that are conserved in all three HBPs. The two histamine ligands are bound with different affinities and exhibit different orientations at their sites. At the high-affinity site, the histamine is perpendicular to the long axis of the b-barrel, while the other ligand at the low-affinity site is oriented parallel to that axis (Figure 10). The histamine at the high-affinity site of HBP2 is in a position comparable to that of heme in the

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Figure 9 Crystal structure of triabin from Triatoma pallidipennis. Ribbon drawings with eight b-strands (blue arrows, labeled A–H) and several helices (red, labeled h1–h3). The N- and Ctermini are marked with white dots. Note the unusual topology of the strands A–C–B–D.

nitrophorins; the low-affinity site is buried in the protein. How the histamines access the two different sites is not clear. To accommodate the hydrophilic histamine, which is positively charged in a physiological environment, both binding sites in the cavity are lined with a number of negatively charged and therefore polar amino acid residues. This is in contrast to typical lipocalins, which usually harbor hydrophobic ligands. In addition to the acidic residues, aromatic side chains contribute to the binding of histamine at the high-affinity site. An interesting variant of a HBP has been recently found in the salivary gland of another tick, Dermacentor reticulatus (Sangamnatdej et al., 2002). cDNA

Figure 10 Crystal structure of the histamine-binding protein HBP2 from Rhipicephalus appendiculatus. Upper and middle panels: ribbon drawings with eight b-strands (blue arrows) and several helical regions (red, the major ones labeled h1 and h2). Lower panel: Ca-backbone structure (light blue) with two histamines bound at the high (H) and low (L) affinity sites. The N- and C-termini are marked with white dots.

Lipocalins and Structurally Related Ligand-Binding Proteins

cloning provided a sequence of 192 amino acids for the mature protein, corresponding to a molecular mass of 22 kDa. The expressed protein exists as a dimer. It is glycosylated, and four putative glycosylation sites are derived from the sequence, which is 36–40% identical with those of the HBPs from R. appendiculatus, described above. Binding experiments with histamine revealed two specific binding sites in the recombinant protein from D. reticulatus. This is in accordance with the HBPs from the other tick, R. appendiculatus. The histamine at one of the sites can be competed by serotonin, which binds with high affinity (Kd of 6  1010 M) to this single site. Thus, this salivary protein is a serotoninhistamine-binding protein (SHBP) with separate binding sites for the two distinct ligands. Though SHBP has not been crystallized, modeling of its sequence into the structure of the HBP2 revealed that the high-affinity site is well conserved, while the low-affinity site is enlarged mainly due to the substitution of an aspartate by glycine. This more spaceous site obviously serves as the binding site for serotonin that is larger than histamine. With respect to the dependence of the synthesis of the HBPs on sex and sex-specific feeding behavior in R. appendiculatus (see above), it is of interest to note that the corresponding SHBP of D. reticulatus is expressed in both sexes. Both types of proteins are produced at increased rates over the feeding period though the two ticks differ in their feeding biology. 4.8.4.5. Developmental Expression of Nitrophorins in Rhodnius prolixus

The four well-studied nitrophorins have been isolated from the salivary glands of the adults of Rhodnius prolixus, where they are stored at different concentrations with NP1 as the dominant isoform, followed by NP2 and NP3, and NP4. This isoform pattern is not constant during the insect’s life cycle, according to recent studies (Moreira et al., 2003). In the first instar, NP2 represents the only major nitrophorin that is accompanied by two recently discovered nitrophorins, named NP5 and NP6, which have been only partially studied so far. At each instar, one additional nitrophorin is acquired, which is NP4 in the second instar and NP1 in the third instar. NP3 appears in the fifth instar and accumulates up to the adult stage. During larval development, the amounts of NP5 and NP6 decrease steadily; they are practically absent in the adult insect. In conclusion, the isoform pattern of the nitrophorins in the salivary gland of R. prolixus is development-specific. It is interesting to note that the nitrophorin form appearing earliest is NP2, the only protein of this group with anticoagulant

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activity in addition to its capability to reversibly bind nitric oxide and histamine. So, this nitrophorin variant provides the first instar nymphs with the full spectrum of activities that are relevant to blood feeding. It should be mentioned that R. prolixus synthesizes in the fat body another heme protein that is released into the hemolymph and taken up by the developing oocytes. This protein with a molecular mass of 15 kDa has a high content of a-helices. Therefore, this Rhodnius heme-binding protein is not a lipocalin. This is further stressed by the lack of any sequence similarity to the nitrophorins (Oliveira et al., 1995; Paiva-Silva et al., 2002). 4.8.4.6. Functions of Nitrophorins and Related Proteins

The main roles of the nitrophorins and functionally related proteins have been described in the context of their crystal structures, which allowed a molecular understanding of their function (see Section 4.8.4.2). So, this paragraph mainly summarizes the functional architectures of proteins adapted to a specific feeding biology. A study of these proteins has revealed a sophisticated molecular concept that enables insects and ticks to feed on vertebrate blood. The overall strategy is to prevent the host’s response to the feeders’ attack in several independent ways that act together in a concerted action. In blood-sucking insects, the nitrophorins act as heme-based stable stores of nitric oxide, which is released into the blood after dilution and as a consequence of the higher pH at the feeding site (Nussenzveig et al., 1995). The nitric oxide is distributed rapidly into the neighboring cells where it binds to the heme of soluble guanylyl cyclase for their activation that finally results in smooth muscle relaxation and vasodilatation. In exchange for nitric oxide, the nitrophorins bind histamine with high affinity that is produced by the host’s mast cells to induce inflammation, immune response, and wound healing. All four nitrophorins studied in detail (NP1 to NP4) exert both activities as nitric oxide donors and as histamine scavengers. Nitrophorin NP2 exhibits anticoagulation activity in addition (Ribeiro et al., 1995). Several other proteins of salivary glands reduce platelet aggregation by binding either ADP (RPAI-1), biogenic amines (ABP), or thrombin (triabin), which all promote blood coagulation. In conclusion, blood-sucking insects have developed a most remarkable battery of lipocalins targeted at diverse steps of the complex blood coagulation cascade in order to maintain the host’s blood flow at the feeding site.

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Ticks have developed a similar strategy to support their blood-feeding by using the same versatile protein fold, that of the lipocalins, to provide binding sites for histamine, but without employing heme, in contrast to the molecular concept of the nitrophorins of bugs. The tick proteins do not transport nitric oxide because they lack heme. The binding of serotonin in addition to histamine in the protein (SHBP) from D. reticulatus is not surprising since biogenic amines in general are established potentiators of blood coagulation and the antiinflammation response, which are also targeted by the biogenic ABP from the bug R. prolixus.

4.8.5. Lipocalins with Unknown Ligands 4.8.5.1. Lipocalins Putatively Related to Development

As described in the foregoing, lipocalins are typically secreted, hence extracellular, proteins found in fluids of insects, like hemolymph (e.g., biliproteins) and saliva (e.g., nitrophorins), although they are also frequently stored as holoproteins within the cells, where they are synthesized (e.g., in the epidermis or fat body) or taken up (e.g., into the eggs). All these lipocalins are typified as carriers of specific ligands that are known, or at least presumed, to serve some metabolic roles in the insects. Well-studied examples of ligands are porphyrin products, nitric oxide, histamine, and biogenic amines. However, a number of proteins have been isolated or cloned from cDNA that were identified as lipocalins on the basis of sequence similarities and the presence of typical motifs in their secondary structures without knowing any real or potential ligands. It is possible that not all lipocalins carry a ligand but may function by, for example, docking to some other proteins as members of a signaling pathway. Under this view, it is interesting to note that several lipocalins without known ligands have been found to be associated with embryonic and postembryonic phases of development, frequently linked to that of the nervous system. 4.8.5.1.1. Lazarillo from Schistocerca americana Studies of the development of the nervous system uncovered a most unusual form of a lipocalin, as it is not free but covalently bound to the outer cell surface. There is no other eukaryotic lipocalin firmly anchored to a membrane (bacterial lipocalins are bound to the outer membrane). Playing a role as a guide to developing neurons, this protein was named Lazarillo after ‘‘Lazarillo de Tomes, a crafty boy who guided a blind man,’’ as explained

by Ganfornina et al. (1995), who discovered this lipocalin by the use of a monoclonal antibody in embryos of the grasshopper Schistocerca americana. A recent overview of Lazarillo has been published by Sa´ nchez et al. (2000a). Lazarillo is highly glycosylated, thus behaving like a 45 kDa protein as affinity-isolated with the monoclonal antibody used to detect this lipocalin. From its cDNA, a molecular mass of 20 kDa was predicted for the mature protein and seven potential N-glycosylation sites, however. The amino acid sequence of Lazarillo is further characterized by hydrophobic sequences at both ends. The N-terminal sequence likely represents a secretion signal, and the C-terminal sequence serves as an anchor for the cell surface attachment via a glycosylphosphatidylinositol (GPI) linkage. As can be seen from Figure 11, the Lazarillo sequence shows four cysteine residues which are most likely oxidized to form disulfide bridges corresponding to the two conserved disulfide bridges in other lipocalins. Moreover, the structurally conserved regions (SCRs) that characterize members of the lipocalin superfamily of proteins (Flower et al., 1993) are well conserved in Lazarillo. At the amino acid sequence level, Lazarillo is about 30% identical with the other lipocalins, a value comparable to other members of this superfamily. A more detailed comparison places Lazarillo in the neighborhood of the biliproteins from P. brassicae and M. sexta, together with various forms of apolipoprotein D from mammals. Based on its relationship to the biliproteins, a homology model for Lazarillo was built using the atomic coordinates of insecticyanin from M. sexta (Sa´ nchez et al., 2000a). This model (Figure 12) shows a close match of the two structures. Remarkably, six of the seven potential glycosylation sites are located on the same side of the protein and close to the C-terminal GPI attachment site, thus resulting in a very polarized glycoprotein. The modeled cavity of Lazarillo also compares well with that of the biliprotein with respect to the distribution of hydrophobic and polar sites. It should be clearly stated at this point that no natural or artificial ligand of Lazarillo is known, though the binding cavity could accommodate a tetrapyrrole, for example. 4.8.5.1.2. Lazarillo-like lipocalins from Drosophila melanogaster The sequencing of the Drosophila genome provided two novel sequences with similarities to Lazarillo. These novel lipocalins were called Drosophila neural Lazarillo (DNLaz) and Drosophila glial Lazarillo (DGLaz), respectively (Sa´ nchez et al., 2000b). As deduced from their sequences, DNLaz is an acidic protein (pI 4.3), while DGLaz

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Figure 11 Alignment of lipocalins putatively related to development. The proteins are gallerin (GAL) from Galleria mellonella, bombyrin (BOM) from Bombyx mori, neural and glial Lazarillo-like proteins (DNL and DGL, respectively) from Drosophila melanogaster, hyphantrin (HYP) from Hyphantria cunea, and Lazarillo (LAZ) from Schistocerca americana. N-terminal secretion signals are boxed in gray. Residues typically conserved in lipocalins are boxed in black. Stars indicate identical residues, double and single points more or less conserved substitutions.

is basic (pI 8.6). Both proteins are 21 kDa in size, exhibit the four conserved cysteines of lipocalins and represent secretory products according to their N-terminal signal sequences. Both may also be glycosylated; there are four potential sites in DNLaz, but only a single one in DGLaz. The two Drosophila proteins are different from other lipocalins and also from each other due to the presence of extra sequences (Figure 11). DGLaz has two of them of different lengths in the middle part of the coding region, which are located in two loops at the bottom of the lipocalin b-barrel. DNLaz shows an extra long C-terminal sequence, which, however, is hydrophilic in contrast to that of Lazarillo. No GPI tail is detected in the two Lazarillo-like proteins from Drosophila.

Figure 12 Homology model of Lazarillo from Schistocerca americana using the atomic coordinates of insecticyanin. The C-terminal helix (green) and the helical region with hydrophobic side chains (orange) might be involved in protein–protein interactions. The seven Asn glycosylation sites are labeled with blue balls. (Courtesy of Dr. Diego Sa´nchez.)

4.8.5.1.3. Gallerin from Galleria mellonella The sequence of gallerin was deduced from a brain cDNA library from the wax moth, Galleria mellonella (Filippov et al., 1995). Gallerin comprises 203 amino acid residues including a signal sequence of 15 residues. According to homology searches, gallerin is related to BBP, the bilin-binding protein

288 Lipocalins and Structurally Related Ligand-Binding Proteins

from P. brassicae, and to the related insecticyanin from M. sexta. (see Section 4.8.3.1). Gallerin shows 40% identity with the two biliproteins, the four cysteines are at comparable positions, and also the three structurally conserved regions (SCRs) of typical lipocalins are present (Figure 11). Unfortunately, gallerin was not compared with the more known neuronal lipocalin Lazarillo, although mentioned in the respective publication (Filippov et al., 1995). The same publication also reports the sequence of another protein, called sericotropin, which was used to pick the gallerin sequence. According to a homology search performed by this author using BLAST, sericotropin is not a lipocalin as claimed by Filippov et al. (1995) since all the reference proteins are in fact of mainly helical structure. This correction is in agreement with the current view of the senior author of this publication (Sehnal, personal communication). 4.8.5.1.4. Bombyrin from Bombyx mori Bombyrin was identified in a brain cDNA library from silkworm, Bombyx mori (Sakai et al., 2001). The sequence of bombyrin was assigned to a lipocalintype protein based on typical signatures like the four cysteines and the structurally conserved regions (Figure 11). A sequence alignment study, performed by this author, revealed that bombyrin is closely related to gallerin (68% identity), while no significant overall homologies (18–23% identity) were found to Lazarillo from the locust and the two Lazarillo-like proteins from the fruit fly. 4.8.5.1.5. Hyphantrin from Hyphantria cunea Another putatively developmental lipocalin, named hyphantrin, was cloned from the fall webworm Hyphantria cunea (Seo and Cheon, 2003). The sequence of hyphantrin was first assumed to represent a BBP though it is not associated with any blue pigment (Seo, personal communication). Only recently has hyphantrin been recognized as a protein with homology to the two Lazarillo-like proteins, to bombyrin, gallerin and Lazarillo (Seo, personal communication) (Figure 11). Hyphantrin is similarly related to various forms of apolipoprotein D and biliproteins (range of 25–30% identity, according to this author). 4.8.5.1.6. Expression and putative functions of developmental lipocalins By the use of the monoclonal antibody that led to the discovery of Lazarillo in S. americana, this protein was not ubiquitious in the nervous system of the embryo but only in subsets of neurons of the central nervous system (CNS), in the enteric nervous system, and in all sensory cells of the peripheral nervous system (Sa´ nchez et al., 1995).

Lazarillo is also expressed in a number of cells at the tips of the Malpighian tubules, in a group of nephrocytes, and in mesodermal cells. Lazarillo is expressed during the entire life cycle from the embryo up to the adult stage. In any case, the occurrence of this lipocalin is restricted to the cell surface, where it seems to be evenly distributed. What role Lazarillo could play in such diverse cell types is presently unclear. Studies of embryos of S. americana suggest that Lazarillo guides axon outgrowth during the development of the nervous system, as axon growth was no longer directed to make the correct contacts in the presence of an antibody against Lazarillo. Similar studies have been performed on the developing brain in the related species, S. gregaria, confirming the essential role of Lazarillo for directed axon pathfinding (Graf et al., 2000). Several hypotheses have been put forward on the mode of action of Lazarillo taking into account typical properties of lipocalins: to be designed as vehicles or receptors of mostly hydrophobic small ligands and to interact with other proteins as a means of intercellular communication (Sa´ nchez et al., 2000a). In the latter speculative role, the large sugar moiety of Lazarillo could play an important role in cellular recognition. In Drosophila, DNLaz and DGLaz are both expressed during embryogenesis in the CNS and in some nonneuronal tissues, but not in the peripheral and enteric nervous systems. The temporal patterns of expression of DNLaz and DGLaz are different during embryogenesis but similar thereafter with a low expression in the larvae and a high expression in pupae and adult flies (Sa´ nchez et al., 2000b). This compares well with Lazarillo that is also mainly expressed in embryos and adults of the locust (Ganfornina et al., 1995). While a hypothesis on the role of Lazarillo in the locust has been formulated on experimental ground, the functions of the Lazarillo-like lipocalins in the fly are completely conjectural. The analysis of loss-of-function mutants, which is now under way, may provide some answer in the near future (Sa´ nchez, personal communication). According to Filippov et al. (1995), gallerin is expressed in the CNS of larvae, pupae, and adults of the wax moth. Gallerin expression was also found in the larval fat body, not in any other tissues. In contrast to the Lazarillo-type proteins and to gallerin, hyphantrin is found only in the pupal stage of H. cunea during a few days (days 4–6). Being mainly expressed in the epidermis, hyphantrin may not represent a typical neuronal lipocalin, although positive Northern plots were obtained with brain and fat body (Seo, personal communication).

Lipocalins and Structurally Related Ligand-Binding Proteins

4.8.5.2. Lipocalins Putatively Related to Reproduction

While the above developmental lipocalins, as they are tentatively grouped here, play only a speculative role in the (growing) insect, i.e., between cells, some other proteins, also belonging to the lipocalin superfamily, seem to be involved in the interactions between the (adult) insect and the environment. As in the developmental forms, ligands are also yet unknown for these lipocalins described below. 4.8.5.2.1. A lipocalin related to sexual behavior A lipocalin apparently involved in sexual behavior has been identified in the cockroach Leucophaea maderae (Korchi et al., 1999). This protein, LmaP22, is specific for adult males, where it is synthesized only in the epidermis of the tergites 2, 3, and 4 and becomes a constituent of the secretion of the dermal gland. The products of this gland are ingested by the female during courtship to act as an aphrodisiac. Partial sequences of the isolated Lma-P22 allowed the preparation of a cDNA encoding a protein with a molecular mass of 19.7 kDa. It consisted of 178 amino acids including an N-terminal signal sequence of 20 residues. Two putative sites for N-glycosylation were identified. Sequence alignments revealed an overall identity of 17–26% between Lma-P22 and several established and putative lipocalins including insecticyanin and gallerin. Moreover, the predicted secondary structure of Lma-P22 was in agreement with the lipocalin family. Whether Lma-P22 carries a ligand under physiological conditions is unknown, though a pheromone or another hydrophobic compound may qualify as candidates. Another protein with high homology to Lma-P22 has also been identified in the tergal secretion of male L. maderae (Cornette et al., 2001). 4.8.5.2.2. A lipocalin related to oviposition A female-specific protein, referred to as Jf23, has been identified in the tarsi of the forelegs of the swallowtail butterfly, Atrophaneura alcinous (Tsuchihara et al., 2000). Jf23 can be extracted from the tarsi as a 23 kDa protein. The sequence of Jf23, as deduced from its cDNA, revealed a preprotein consisting of 203 amino acid residues, including a leader peptide of 15 amino acids. The calculated molecular mass of 20 296 Da of the Jf23 precursor is in good agreement with the estimated size of the isolated protein. Nearly half of the amino acids are hydrophobic. With a sequence identity of 38%, Jf23 is most closely related to BBP, the bilin-binding protein from P. brassicae, followed

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by insecticyanin, gallerin, and some mammalian lipocalins. A lipocalin structure of Jf23 is further supported by the presence of the conserved cysteines and of two structurally conserved regions at the corresponding locations. The presence of Jf23 in the female tarsi suggests that it may play a role in the transduction of plant chemical signals in the context of egg deposition. In fact, in electrophysiological experiments with the sensilla of female tarsi the response to compounds of a host plant was partially suppressed after pretreatment of the sensilla with antiserum against Jf23.

4.8.6. Fatty Acid-Binding Proteins 4.8.6.1. Variation of the Lipocalin Structure

Fatty acid-binding proteins (FABPs) are ubiquitous in animals. Members of this superfamily have been isolated from numerous vertebrates including humans, and from a variety of tissues such as intestine, liver, and heart muscle. While lipocalins are said to function typically as transporters of a variety of lipophilic molecules between cells, i.e., in the extracellular space (e.g., biliproteins in insect hemolymph or retinol-binding protein in vertebrate serum), FABPs are intracellular proteins and specialized as an almost single class of ligands, as their name indicates. FABPs are acidic proteins (pI 5) and smaller than lipocalins, being composed of about 130 amino acid residues, corresponding to 15 kDa proteins. On the other hand, FABPs share the overall structure of a b-barrel and a repeated þ1 topology with the lipocalins. Because of some differences in the construction of the b-barrel, however, they are grouped as a separate superfamily within the calycin structural superfamily of proteins. Two other groups of intracellular proteins exhibit the same structural motif as the FABPs, which therefore all belong to the same superfamily. These proteins are the cellular retinol- and retinoic acid-binding proteins (CRBPs and CRABPs), respectively, reviewed by Banaszak et al. (1994) and Newcomer (1995). Only one representative has been identified in an insect to date (see Section 4.8.6.3). Because of their location in distinct compartments, the FABPs/CRBPs/CRABPs and the lipocalins are also referred to as intracellular and extracellular lipid-binding proteins, iLBPs and eLBPs, respectively. As a further difference between the two types of proteins, the orientation of the ligand is different. In the iLBPs, the polar group of the ligand (e.g., the carboxyl group of a fatty acid) is directed to the interior of the cavity, while it points to the barrel opening in the eLBPs (e.g., the carboxyl groups of

290 Lipocalins and Structurally Related Ligand-Binding Proteins

the tetrapyrroles). The crystal structure of the first representative of the FABPs was described in 1988 for the P2 myelin protein from the vertebrate peripheral nervous system (Jones et al., 1988). To date, the crystal structures of only two insects FABPs have been resolved, as described below. General overviews of the two major forms of b-barrel ligandbinding proteins, the lipocalins and the FABPs, have been written by Flower et al. (1993) and LaLonde et al. (1994), while the excellent review by Banaszak et al. (1994) is much more comprehensive and detailed. Overall, the b-barrel of the FABPs is flatter and more clamlike in comparison to the more spherical lipocalin barrel. The major difference in the folding pattern is that the b-barrel of the FABPs is constructed not by eight, but by 10 antiparallel b-strands that are hydrogen-bonded to form two sheets of four and six strands, respectively. Like in the lipocalins, the strands forming the two sheets are in orthogonal orientation to one another. A characteristic feature of the general folding of the FABPs is the gap between the fourth and the fifth strand, assigned bD and bE. The first two strands are separated by a helix–turn–helix motif that is not present in the lipocalins. A scheme of the sequential arragement of secondary structures of the FABPs is depicted in Figure 13. The helical parts of the N-terminal sequence occlude the b-barrel similarly to the first loop, the large O loop, in the lipocalins (see Section 4.8.2). FABPs lack the C-terminal helix of the lipocalins. The cavity of the b-barrel is mostly lined with hydrophobic side chains, but polar groups are also present. It is considerably more spacious than is required to bind a single molecule

of fatty acid. In contrast to the situation in the lipocalins, it is unknown where or how the ligand enters the cavity of a FABP. It is speculated that the loading and unloading of the ligand may require dynamic changes in the protein folding that could be induced by the fatty acid itself. Nuclear magnetic resonance (NMR) studies at least indicate some flexibility in the region of the helices possibly serving as a portal for the ligand. The gap between the strands bD and bE is apparently not suitable for this role. Despite the widespread occurrence of FABPs, only two representatives from insects have been purified, crystallized, and studied for their structure to date. These FABPs have been isolated from M. sexta and S. gregaria. Several other proteins have been identified as FABPs on the basis of sequence similarity with established FABPs, as described below (see Section 4.8.6.3). A number of FABPs from insects and mites are well-known allergens, as will be discussed separately (see Section 4.8.7). Those readers with particular interest in muscle-type FABPs from mammals and insects may consult the comparative article by Zanotti (1999). More comprehensive reviews of the FABPs from vertebrates are from Veerkamp and Maatman (1995) and Glatz and van der Vusse (1996). For more details and an overall view of lipid transport in insects, see Chapter 4.6. 4.8.6.2. Fatty Acid-Binding Proteins with Known Crystal Structure

4.8.6.2.1. Fatty acid-binding proteins from Manduca sexta Two FABPs, referred to as MFB1 and MFB2, have been isolated from the midgut cytosol of fifth instar larvae of M. sexta, as the most

Figure 13 Aligment of fatty acid-/retinoic acid-binding proteins. The proteins are the fatty acid-binding proteins from Schistocera gregaria (Sg), Manduca sexta (Ms2: MFB2), Blo t 13 from Blomia tropicalis (Bt13), Lep d 13 (Ld13) from Lepidoglyphus destructor, and the cellular retinoic acid-binding protein (CRABP) from M. sexta. Black boxes show residues typically conserved in fatty acid-binding proteins. The residues boxed in gray (Gly in Sg and Leu in Ms2) are apparently responsible for the conformation of the bound fatty acid (see text for details). Below the sequences, the approximate positions of helices and b-strands are labeled as red and blue bars, respectively. The gap between the strands D and E is marked in yellow. Asterisks indicate identical residues, double and single points more or less conserved substitutions.

Lipocalins and Structurally Related Ligand-Binding Proteins

abundant proteins amounting to 2% and 12%, respectively, of the total soluble protein (Smith et al., 1992). The predominant form, MFB2, has been crystallized and its structure determined as the first representative of an insect FABP (Benning et al., 1992). The crystal structure of MFB2 (Figure 14) agrees with the folding motif of vertebrate FABPs. The b-barrel is made up of 10 up-and-down b-strands that are combined to form one sheet with four and one sheet with six strands. The N-terminal sequence is characterized by two helices that link strand A and strand B with a helix–turn–helix motif giving rise to the conserved clamlike overall structure of the barrel. The amino acid side chains defining the cavity

Figure 14 Crystal structure of the fatty acid-binding protein MFB2 from Manduca sexta. Ribbon drawings with ten b-strands (blue arrows, labeled A–J) and two helices (red) located between strands A and B (cf. Figure 13). The N- and C-termini are marked with white dots. The bound fatty acid is shown in spacefilling representation with the carboxyl oxygens in red. Note the gap between strands D and E.

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for binding a fatty acid are hydrophobic, and most of them are uncharged. The aliphatic chain of the ligand is in contact with these hydrophobic sites, while the polar carboxyl group is hydrogen-bonded mainly to a conserved arginine (Arg127) and a tyrosine (Tyr129) located close to the C-termius that lacks the lipocalin-typical helix. The carboxyl group of the ligand is additionally bound to a sulfate which may be an artifact since the protein was crystallized from an ammonium sulfate solution. It is unknown whether a polar small molecule plays a role in the binding of a fatty acid in a physiological environment. While the position of the carboxyl group of the fatty acid in MFB2 is comparable to that in the vertebrate proteins, the conformation of the aliphatic chain is different. This is obviously due to a difference in the amino acid sequences which has an impact on size and shape of the cavity. In MFB2, a bulky leucine (Leu32) residue, corresponding to a glycine or alanine in the vertebrate proteins, forces the aliphatic tail of the fatty acid beyond C6 to an opposite orientation compared to the vertebrate proteins. The minor form, MFB1, of the FABPs from the midgut of M. sexta has not been crystallized but its three-dimensional structure is thought to be very similar to that of MFB2. As derived from their cDNA sequences, MFB1 and MFB2 are of identical length comprising 131 amino acid residues, corresponding to molecular masses of 14 834 Da and 14 081 Da, respectively. As in most other FABPs, the N-termini are blocked also in the two Manduca proteins. The sequence identity between MFB1 and MFB2 is 56%, which is low for proteins of the same type and from the same tissue and species. The identities of the FABPs from the moth to those from various vertebrates are around 30% or lower. Remarkably, the two moth FABPs are not evenly distributed along the midgut. MFB1 is more concentrated in the anterior part but also present in the posterior part, while MFB2 is limited the posterior part of the midgut. Neither of the two proteins could be detected in extracts from fat body, muscles, and eggs. Like the lipocalins, MFB1 and MFB2 bind ligands in a 1 : 1 molar ratio. As isolated from the midgut, both proteins were loaded with a mixture of saturated and unsaturated fatty acids among which C16:0 and C18:2 fatty acids were predominant. The bound fatty acid can be easily exchanged in MFB1, but not in MFB2. Saturation binding experiments performed with MFB1 and radiolabeled oleic acid yielded an apparent dissociation constant of 14 mM for the exchange reaction. This is, of course, not a true measure for the affinity of the ligand to the apoprotein.

292 Lipocalins and Structurally Related Ligand-Binding Proteins

4.8.6.2.2. Fatty acid-binding protein from Schistocerca gregaria The first invertebrate FABP was isolated from S. gregaria. It was discovered in extracts from the flight muscles from adult locusts where it makes up about 18% of the total cytosolic protein (Haunerland and Chrisholm, 1990; Haunerland et al., 1993). As a soluble acidic protein (pI 5.2) with a molecular mass of 15 kDa and fatty acids bound in a 1 : 1 molar ratio, this locust protein fits quite well the overall characteristics of other FABPs (see Section 4.8.6.1). By contrast, however, the Nterminus of the locust FABP was not blocked. The profile of the fatty acids, bound to the FABP from S. gregaria as isolated, was very similar to that of the two proteins from M. sexta. The locust FABP crystallized as a dimer in which the monomers are connected via the conserved helix–turn–helix motif that occludes the barrel like a lid (Haunerland et al., 1994). Overall, the protein shows the same conserved tertiary structure that is characteristic of members of this superfamily (Figure 15). These features are the highly conserved b-barrel with 10 antiparallel up-and-down b-strands with a gap between the bD and bE strands (Figure 15). The interior of the barrel contains 23 ordered water molecules, five of which are conserved between the proteins from S. gregaria and M. sexta. The locust FABP was obviously studied in the apoprotein form, as no fatty acid-ligand was found in the cavity suggesting its loss during the purification of the protein. However, the conserved C-terminal arginine and tyrosine residues that provide binding sites for the carboxyl group of the fatty acid in the moth MFB2, are also present in the locust FABP. While this suggests a similar mode of ligand binding, the conformation of the hydrocarbon tail of the ligand in the locust protein is expected to be opposite to that in MFB2, as the relevant bulky leucine (Leu32) in MFB2 corresponds to a small glycine (Gly34) in the locust FABP. This ligand conformation would be comparable to that in the vertebrate FABPs. The FABP from S. gregaria, which comprises 133 amino acids (Figure 13), shows unexpected low similarity to the two proteins from M. sexta and to those from vertebrates (about 30% sequence identity), while it is closer related (41% identity) to the FABP from human cardiac muscle (Price et al., 1992). Generally, FABPs from the same type of tissue but from different species turned out to be more similar to each other than those from different tissues in the same species. A comparison between the FABPs from insect flight muscles and midgut, for example, would be very interesting in this respect. Not surprisingly, however, the amino acid sequences of the FABPs from the flight muscles of the two

Figure 15 Crystal structure of the fatty acid-binding protein from Schistocerca gregaria. Upper panel: ribbon drawing showing ten b-strands (blue arrows, labeled A–J) and two helices (red) located between strands A and B (cf. Figure 13). The N- and Ctermini are marked with white dots. Lower panel: Ca-backbone structure of the upper representation flipped around a vertical axis. The protein was isolated in the apo form.

locusts S. gregaria and Locusta migratoria are nearly identical (see Section 4.8.6.2.3). The FABP from S. gregaria has also been studied at the gene level (Wu and Haunerland, 2001; Wu et al., 2001). In mammals, the FABP genes are characterized by three introns of varied length inserted at identical positions of the coding sequence. The locust gene shows only two introns, exactly corresponding to the first and third mammalian introns. This suggests a common evolutionary orgin of the FABPs. The promoter region contains a palindrome with a unique sequence of 19 bp that is essential for the induced expression of the gene by fatty acids. How this transcriptional effect is

Lipocalins and Structurally Related Ligand-Binding Proteins

mediated is unknown. It has been hypothesized that the conserved helix–turn–helix motif of a FABP could play a role in gene regulation, as it is reminiscent of a known transcription factor motif. 4.8.6.2.3. Fatty acid-binding protein from Locusta migratoria Another locust FABP is known from the flight muscles of L. migratoria (Maatman et al., 1994). Cloning and heterologous expression of its cDNA has provided data on its sequence as well as binding of various ligands to the recombinant FABP. Although crystals of the protein were obtained, structural data have not yet been published. The L. migratoria FABP comprises 133 amino acids, corresponding to a molecular mass of 14 935 Da, and has a nonacetylated N-terminus, like in the other locust FABP. The sequence of the FABP from L. migratoria shows 98% identity with that of the S. gregaria homolog and 42% identity to that of the human muscle protein (Maatman et al., 1994). According to the conserved features of its primary structure, the FABP from L. migratoria is expected to also posses the conserved tertiary structure known for this protein superfamily. Binding experiments, using the recombinant protein, provided a Kd for oleic acid of 0.5 mM and confirmed the 1 : 1 stoichiometry of ligand binding. The FABP from L. migratoria is less acidic (pI 6.1) than that from S. gregaria (pI 5.2). 4.8.6.3. Fatty Acid-Binding Proteins with Unknown Crystal Structure

The sequence of a FABP from the moth Heliothis zea, the corn earworm, has been deposited in the databases (Heilmann, 1998). It has been identified on the basis of homology of its cDNA-derived sequence with that of established members of the FABP superfamily. Sequence alignment studies with insect FABPs, performed by this author using the Clustal W program, revealed a relatively high identity of the muscle protein from H. zea with the two midgut proteins from M. sexta (44% and 39% for MFB1 and MFB2, respectively), while the corresponding values for almost all other FABPs, including those from flight muscles of the two locusts, were in the 20% range. Another FABP has been reported (as a DNA sequence fragment) from the honeybee, Apis mellifera, where it is expressed only in larvae of workers, not of queens (Evans and Wheeler, 1999). A 14-kDa protein with a free N-terminus was purified from the flight muscles of the bug Dipetalogaster maximus, and identified as a heart-type FABP based on its N-terminal sequence (Cavagnari et al., 2000). The sequencing of the genome of Drosophila melanogaster has revealed

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several genes and gene fragments obviously coding for FABPs. Similarly, one gene coding for a FABP of 141 amino acids and a calculated molecular mass of 15 071 Da has been cloned from the mosquito Anopheles gambiae (Favia et al., 1996). There are other FABPs that are discussed subsequently as they have been identified as important allergens, a property also shared by a number of lipocalins (see Section 4.8.7). The FABPs are not the only intracellular ligandor lipid-binding proteins characterized by a b-barrel of ten antiparallel strands. The same folding-type is also found in a family of proteins called cellular retinol- and retinoic acid-binding proteins (CRBPs and CRABPs), respectively, as mentioned above (see Section 4.8.6.1). Their presence in vertebrates is well known, and the crystal structures of several forms have been described (reviews: Banaszak et al., 1994; Newcomer, 1995). Only one insect member of the CRABP family has been identified to date. This protein isolated from M. sexta exhibits 71% identity with bovine and murine CRABP I and could therefore well be modeled into the crystal structures of the vertebrate homologs (Mansfield et al., 1998). The sequence of this moth protein matches well the conserved FABP pattern (Figure 13). The natural ligand is unknown but the binding cavity could accommodate retinoic acid, for example. 4.8.6.4. Functions and Developmental Expression of Fatty Acid-Binding Proteins

As their name indicates, FABPs are intracellular transporters specialized for fatty acids as ligands. Though it seems evident that fatty acids require a carrier system that is mobile in the aqueous cytosol while offering a suitable site for the hydrophobic ligand, the implicated uptake and release of fatty acids by these binding proteins in a physiological environment has not yet been demonstrated clearly (Haunerland, 1997). As discussed above (see Section 4.8.6.1), the conserved folding structure of the FABPs does not suggest an opening of the cavity through which the fatty acid could access or exit the protein cavity. This is quite in contrast to the lipocalins with their calyx structure (see Section 4.8.2). While experiments in vitro with the M. sexta protein MFB1 demonstrated that a bound ligand can readily be exchanged, this was not possible in the case of the homolog MFB2 (Smith et al., 1992). Hence, there may be significant differences between the various FABPs with respect to the ease and the rate of ligand exchange, as well as to the affinities for potential ligands. It is presumed that these different binding properties can be traced back to differences in the amino acid sequences translating

294 Lipocalins and Structurally Related Ligand-Binding Proteins

into differences in the ligand-binding pocket of the proteins. It would be of interest to compare the two FABPs from M. sexta in this respect and to relate their structural differences to the physiology of those parts of the midgut, where they are predominantly localized. The same considerations may be applied to the various tissue-specific isoforms of FABPs that are known from vertebrates. In spite of the lack of understanding of the mechanisms of ligand uptake and release in the FABPs at the level of their crystal structure, these proteins are apparently tightly linked to lipid metabolism, specifically to the utilization of fatty acids for energy production (Van der Horst et al., 1993). This seems to hold for insects as well as vertebrates. In the latter, the heart muscle is a typical tissue specialized on lipid oxidation to generate energy, and it contains a high concentration of tissue-specific FABP. In insects, migratory species similarly depend on the use of fatty acids to fuel long-distance flight. The flight muscles of locusts, for example, are an exceptionally rich source of FABP (Haunerland et al., 1993). Under the conditions of extended flight, lipids are released from the fat body and transported via specialized hemolymph proteins to the flight muscles (see Section 4.8.8.2), where the fatty acids are taken up for b-oxidation (reviews: Haunerland, 1997; Ryan and Van der Horst, 2000) (see Chapter 4.6). The FABPs are supposed to be responsible for the intracellular transport of the fatty acids from the cell membrane to the mitochondria. It has been proposed that FABPs may also play a role in regulating the intracellular concentration of free fatty acids to prevent damage due to their detergent effect. In S. gregaria, FABP was localized in the cytosol as well as in the nucleus but not in mitochondria. The nuclear presence might suggest a genomic action, possibly on the expression of the FABP gene itself, as the transcription of the FABP gene can be enhanced by fatty acids. Thus, this transporter protein could be regulated according to physiological needs (Haunerland et al., 1992a; Chen and Haunerland, 1994). The FABP from S. gregaria flight muscle is present only in the adult. Only mature locusts are able to perform extended flight that requires the presence of this type of protein. The concentration of FABP starts to rise a few days after adult ecdysis reaching a plateau after about 10 days. This increase in FABP protein follows a transient but strong increase in its mRNA before it returns to a constant, low level (Haunerland et al., 1992a; Haunerland, 1997). The high capacity to utilize fatty acids for energy production develops only several days after the molt

to the adult stage, in parallel with the ability to perform extended flight. Overall, FABP synthesis appears to be part of the developmental program of an adult-specific muscle. This is also demonstrated by the application of inhibitors of metamorphosis that prevent the development of mature flight muscles and, in consequence, the appearance of the FABPs (Haunerland et al., 1993).

4.8.7. Lipocalins and Fatty Acid-Binding Proteins as Allergens Proteins with a b-barrel structure are of growing medical interest after several major respiratory allergens of mammalian and insect origin were identified as lipocalins and fatty acid-binding proteins, respectively (reviews: Ma¨ ntyja¨ rvi et al., 2000; Arlian, 2002). Though these calycins share the conserved protein architecture, conserved sequence regions, and some overall sequence homology, the structural determinants of their allergenicity have not yet been identified. It is evident, however, that it is not the b-barrel per se since by far not all lipocalins induce an allergic immune response. Well-studied examples of allergenic lipocalins from vertebrates are Bos d 2 from cow dander and Mus m 1 from mouse urine. The major sources for arthropod allergens from the superfamilies of lipocalins and fatty acid-binding proteins (FABPs), respectively, are species belonging to cockroaches, bugs, and mites. Among the numerous allergens from cockroaches, Bla g 4 from Blattella germanica has been studied at the structural level and recognized as a typical lipocalin (Arruda et al., 1995). The sequence of Bla g 4 comprises 182 amino acids with a calculated molecular mass of 20 904 Da, as deduced from its cDNA. The N-terminal 12 residues suggest that the encoded protein is secreted. The sequence contains one potential N-glycosylation site. The allergenic nature of the protein was confirmed using its recombinant form. A gene coding for Bla g 4 was also identified in Periplaneta americana, but its expression could not be demonstrated in this species, however. Searches for similarity to the Bla g 4 sequence revealed several established lipocalins from insects and vertebrates as related proteins. This cockroach allergen contains all three structurally conserved regions (SCRs) and the four conserved cysteines that define membership in the lipocalin family (Figure 8). Though the overall identity is low, as typically found among lipocalins, the sequence of Bla g 4 was successfully modeled into the crystal structure of the bilin-binding protein (BBP) from P. brassicae. The biological function of

Lipocalins and Structurally Related Ligand-Binding Proteins

Bla g 4 is unknown as is the binding of any ligand, although a role as a carrier of pheromones has been proposed. Another group of insects with high allergenic potency are hematophagous bugs. The allergenic activity is stored in the salivary glands in many Triatoma species. In T. protracta, most of this activity is associated with a 19 kDa protein, named procalin, that comprises, as predicted from its cDNA, 169 amino acids with an N-terminal hydrophobic signal sequence (Paddock et al., 2001). Unlike Bla g 4, procalin has no potential glycosylation site. An analysis of the procalin sequence predicts that procalin adopts a typical lipocalin fold with eight bstrands and a C-terminal helix. In fact, the closest homologs of procalin are the salivary platelet aggregation inhibitors from R. prolixus and triabin from T. pallidipennis with its unusual lipocalin fold (see Section 4.8.4.3). Moreover, the similarity of procalin to several established lipocalins, such as the moth biliproteins, retinol-binding protein, and b-lactoglobulin, has already been noted. One of the allergens of the dust mite Blomia tropicalis, a most prevalent species in tropical areas, shows 40% sequence identity with several FABPs from vertebrates and a flatworm. In fact, this allergen with the official name Blo t 13 represents a typical FABP with respect to size (130 amino acids; molecular mass of 14 800 Da) and binding specificity for fatty acids. Moreover, Blo t 13 is characterized by ten up-and-down b-strands, and its sequence could well be fitted into the conserved crystal structure of FABPs (Caraballo et al., 1997; Puerta et al., 1999). Further FABP allergens from mites are Lep d 13 from Lepidoglyphus destructor and Aca s 13 from Acarus siro (Eriksson et al., 1999, 2001). The sequences of Blo t 13 and Lep d 13 are presented in Figure 13 in comparison to those of other FABPs.

4.8.8. Ligand-Binding Proteins That Are Not b-Barrels 4.8.8.1. Odorant-Binding Proteins

Olfaction plays a major role in the lifes of insects and vertebrates (see Chapter 3.15). It is based on air-borne chemicals that could be pheromones (see Chapter 3.14), plant-derived compounds or other signaling substances. Volatile compounds are usually small and hydrophobic, requiring a carrier system that takes them from the cuticular surface of insects or from the nasal epithelium of vertebrates to the olfactory receptors. This carrier function is supposed to rely on a specialized set of proteins, called odorant-binding proteins (OBPs). They comprise

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the general odorant-binding proteins that occur in both sexes and have no clear ligand preferences and the pheromone(sex attractant)-binding proteins, which are male-specific and specialized in binding a defined ligand, the female pheromone. Many of these OBPs from insects and vertebrates have been purified, cloned, and studied for their functional properties (Pelosi and Maida, 1995; Field et al., 2000). OBPs from insects and vertebrates share several features: they are small (15 to 20 kDa), acidic (pI 5) and secreted in high concentrations. However, there is no sequence similarity between insect and vertebrate OBPs. This is reflected in their three-dimensional structures: the OBPs from vertebrates have been clearly identified as members of the lipocalin superfamily with all the features of b-barrel proteins, while those from insects are rich in a-helical structures and therefore not lipocalins. The protein folds of two OBPs from insects have recently been elucidated by NMR spectroscopy and X-ray diffraction, respectively: these are a hemolymph protein from the beetle Tenebrio molitor (Rothemund et al., 1999) and a pheromonebinding protein in the complex with bombykol from Bombyx mori (Sandler et al., 2000). Both studies revealed a new structural class of proteins defined by six a-helices forming a small hydrophobic core that provides a ligand-binding site. Three disulfide bridges are also conserved. Recently, an X-ray study of a chemosensory protein from the moth Mamestra brassicae demonstrated another novel fold consisting of six a-helices linked by a–a loops (Lartigue et al., 2002). Evidence for a predominant helical structure has also been obtained for a chemosensory protein from the honeybee, studied by CD spectroscopy (Briand et al., 2002). Furthermore, the genome of Drosophila has been searched for OBP genes; all identified 38 genes clearly featured a six-helical fold (Graham and Davies, 2002). It is remarkable from an evolutionary standpoint that insects and vertebrates developed completely different protein structures to achieve functionally equivalent tools for chemical communication. 4.8.8.2. Lipophorins

Hydrophobic molecules in general need amphiphilic transport systems operating in the extracellular space between the tissues as well as in the intracellular milieu (see Chapter 4.6). This aspect has already been discussed in the intracellular transport of fatty acids by a specialized set of binding proteins, the fatty acid-binding proteins (FABPs), which represent a variation of the b-barrel structure as it is found in the lipocalins (see Section 4.8.6.1). Insects performing extended flight like migratory

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locusts and sphingid moths rely on the oxidation of lipids to produce energy. The lipids are stored in the fat body and activated to fuel flight by the conversion to diacylglycerols, representing their transport form. The transport of lipids to the cells, which finally b-oxidize the fatty acids, is performed by a special carrier system, called lipophorin. Lipophorin represents a large lipoprotein complex composed of two integral units of different size, apolipophorin I and apolipophorin II, with molecular masses of 250 kDa and 80 kDa, respectively. Lipophorin is charged with a variable amount of lipid depending on the intensity of lipid utilization by the insect and can also be associated with steroids, carotenoids, or other hydrophobic compounds. Another protein reversibly associates with the core lipophorin to stabilize this carrier complex and to allow more diacylglycerol to bind. This additional protein, apolipophorin III, with a molecular mass of 19 kDa is highly abundant in the hemolymph as a lipid-free monomer, according to studies in L. migratoria and M. sexta. While the amino acid sequences of apolipophorin III are known from a number of insects, the only available crystal structure is that of the protein from L. migratoria (Breiter et al., 1991). The locust apolipophorin III does not represent a lipocalin or a structural variant thereof like the fatty acid-binding proteins (FABPs). By contrast, it consists of five long a-helices connected by short loops. The helices show a clear amphiphilic arrangement of hydrophobic and hydrophilic amino acid residues, thus fitting the physicochemical requirements to interface the lipid and aqueous compartments. The sequence homology between the apolipophorins III from L. migratoria and M. sexta suggests a fairly conserved structure of this lipophorin partner. For more details on lipophorin in the overall context of lipid transport (see Chapter 4.6).

4.8.9. Crustacean Lipocalins with Established Crystal Structure Lobster (Homarus gammarus) is of long-standing interest not only for gourmets but also for scientists interested in the secrets of the blue coloration of its shell turning into appetizing red upon cooking. The red color is due to astaxanthin (3,30 -dihydroxyb,b-carotene-4,40 -dione), a carotenoid metabolically derived from b-carotene by oxidation at both cyclohexene end rings to produce an extended chromophore with a light absorption maximum in the range of 470–490 nm, depending on the solvent (review: Kayser, 1985). In lobster, astaxanthin is noncovalently bound to protein in a complex, called crustacyanin, with an absorption maximum of the

carotenoid at 630 nm, resulting in a blue protein. This huge bathochromic shift of more than 100 nm upon binding is due to a specific arrangement of the red carotenoid in the native protein complex (i.e., before cooking), as described below. Crustacyanin or, more specifically, a-crustacyanin is a multimeric protein complex, which comprises eight heterodimeric units, called b-crustacyanins. Each of the b-crustacyanin dimers is composed of two types of monomers (the known monomers are A1–3 and C1–2) of similar size (20 kDa) each with one bound molecule of astaxanthin. Thus, the holoprotein with a molecular mass of 320 kDa contains 16 molecules of astaxanthin. The amino acid sequences of the 20 kDa subunit show homology to members of the lipocalin family (e.g., retinolbinding protein; biliproteins from P. brassicae and M. sexta; see Section 4.8.3.1), and the predicted tertiary structure is consistent with the antiparallel b-barrel fold. Moreover, two of the sequence regions and the two disulfide bridges that are conserved in the lipocalins are also present in the crustacyanin subunits. On the other side, the homology between subunits may be surprisingly low (for example, 85 kDa) proteins that achieve full catalytic activity in the presence of mM calcium concentrations. Some of the cPLA2s are of particular interest because they show a marked preference for PL substrate with AA in the sn-2 position. These enzymes are thought to mediate the first step in eicosanoid biosynthesis. cPLA2s have been purified from several mammalian sources (Dennis, 1994, 1997; Balsinde et al., 1999), including rat mesangial cells, human monocytes, human platelets, leukocytes, Swiss mouse 3T3 cells, glomerular mesangial cells, mouse keratinocytes, and mouse peritoneal macrophages. These enzymes are

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variously stimulated by proinflammatory cytokines, tumor necrosis factor, lipopolysaccharides (LPSs), and mitogens. The stimulations result in the translocation of the enzyme from the cytosol to the cellular membranes, where AA is selectively released from PLs. While the concentration of free AA is maintained at submicromolar levels by a reacylation pathway, stimulation of PLA2 activity produces a rapid increase in free AA, which can then be available for eicosanoid biosynthesis. The cPLA2s are upregulated by several mechanisms. One is a calcium-dependent translocation to the membrane fractions of cells, as just mentioned. Alternatively, the enzymes are activated and deactivated inphosphorylation /dephosphorylation cycles. In other cases, a receptor-associated G protein activates cPLA2s. Finally, some of these enzymes are activated by transcriptional activation, resulting in increased levels of cPLA2 protein. The key point is that cPLA2s represent the first step in eicosanoid biosynthesis. Figure 1 provides an overview of the major eicosanoid biosynthetic pathways. The cyclooxygenase (COX) pathways yield the PGs and thromboxanes and the LOX pathways convert AA into various hydroperoxyeicosatetranoic acids (HPETEs) and hydroxyeicosatetranoic acids (HETEs). These species are themselves biologically active; they are also potential substrates for further metabolism to leukotrienes and still other biologically active products. The epoxygenases are cytochromes P450 (see Chapter 4.1), which yield the various epoxyeicosatrienoic acids. 4.9.2.2.2. The cyclooxygenase pathways PGs are C20 carboxylic acids with a five-membered ring variously substituted at C9 and C11, and two aliphatic chains featuring a substitution at C15 and one, two, or three double bonds. Three PUFAs, C20:3n
6, C20:4n
6, and C20:5n
3, are potential substrates for the COX pathways, although AA is the commonest substrate among mammals. The PGs always have two fewer double bonds than their parental PUFAs, giving rise to the 1-, 2-, and 3-series PGs (Figure 2). PGs are defined by the substitutions at C9 and C11. PGE, for example, features a keto function at C9 and a hydroxyl function at C11. Specific PGs are identified by combining the number and letter designations. PGE1, to continue the example, features one double bond at C13. The parental fatty acid is C20:3n
6. Figure 3 indicates that PG biosynthesis requires three enzyme steps. First is the COX step, which catalyzes the bis-oxygenation of AA to form the endoperoxide PGG2. Second, the endoperoxide

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Figure 1 A sketch of arachidonic acid (AA) metabolism based on the biomedical background. Three fatty acids, C20:3n
6, AA, and C20:5n
3 are potential substrates for eicosanoid biosynthesis. Chemical structures are denoted by numerals: 1, cellular phospholipid; 2, hydrolyzed AA; 3, prostaglandin E2; 4, 5-hydroperoxyeicosatetraenoic acid; 5, leukotriene B4; 6, 11,12-epoxyeicosatrienoic acid; 7, lipoxin A. Capital letters indicate major enzyme systems responsible for eicosanoid biosynthesis: A, phospholipase A2; B, cyclooxygenase (COX); C, cytochrome P450 epoxygenase; D, lipoxygenase (LOX). (Reproduced with permission from Stanley, D.W., 2000. Eicosanoids in Invertebrate Signal Transduction Systems. Princeton University Press, Princeton, NJ; ß Princeton University Press.)

undergoes a two-electron reduction at C15, catalyzed by a peroxidase activity. The peroxidase activity yields PGH2, from which other biologically active PGs are produced. The COX and peroxidase activities are juxtaposed in a single protein, known for many years as PG endoperoxide synthase, or PGH synthase. It is now termed COX. A couple of interesting features of COX help us understand PG biosynthesis. For one, the COX undergoes ‘‘suicide’’ inactivation, thought to be an intrinsic property of the enzyme. This was first recognized because COX is inactivated before all available substrate is converted into product, in

mammalian cells after about 1300–1400 catalytic operations (Smith and Marnett, 1991; Smith et al., 1991). The suicide step is a feature of the COX component of the protein because the peroxidase activity remains after the COX has faded. Suicide inactivation may set an upper limit on cellular capacity for PG biosynthesis. A second interesting feature of mammalian COXs is the intracellular localization of the protein. COX is a glycoprotein associated with membrane fractions, mainly endoplasmic reticulum and to some extent the nuclear membrane. The enzyme was originally thought to feature transmembrane domains,

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Figure 2 Cyclooxygenase (COX) converts certain C20 PUFAs into their respective 1-, 2-, and 3-series prostaglandin (PG) products. Each PG series features two fewer double bonds than its corresponding precursor PUFA. 1, C20:3n
6; 2, PGE1; 3, C20:4n
6; 4, PGE2; 5, C20:5n
3; 6, PGE3. The lower panel presents the ring features of five prostaglandins, where R represents the aliphatic chains that appear on the complete structures. (Reproduced with permission from Stanley, D.W., 2000. Eicosanoids in Invertebrate Signal Transduction Systems. Princeton University Press, Princeton, NJ; ß Princeton University Press.)

but current thinking places the enzyme entirely within the lumen of the endoplasmic reticulum and nuclear membranes (Otto and Smith, 1995). This model held sway until early 1991 when it became clear that some cells express another form of COX. The two forms are now called COX-1 and COX-2. All the preceding remarks are based on our understanding of COX-1, which is thought to serve as a housekeeping isozyme. That is, it is responsible for biosynthesizing PGs active in physiological homeostasis. This enzyme is constitutively expressed in most mammalian tissues, although not in all cells within a tissue. COX-1 is thought to release PGH2 into the cytosol where it is converted by other enzymes into biologically active PGs. The active PGs may act within the cell or may exit the cells, probably assisted by a PG transporter (Kanai et al., 1995). COX-2 mediates the biosynthesis of PGs for inflammatory processes, ovulation, and mitogenesis. In contrast to COX-1, COX-2 is not expressed in most mammalian cells (Smith et al., 1996). However, it can be induced rapidly in many cells, including fibroblasts, endothelial cells, monocytes, and ovarian follicles. COX-2 is an inducible, rather

than constitutive enzyme, and its expression is increased tremendously, from 10- to 80-fold by proinflammatory or mitogenetic factors, such as cytokines and tumor-promoting phorbol esters. While also associated with the membrane fractions of cells, COX-2 is mainly associated with the luminal surface of nuclear membranes. It may release PGH2 into the nucleus, and the PGH2 or another PG derived from it may interact with nuclear proteins to influence gene expression. Hence, COX-1 and COX-2 may represent two unrelated pools of active enzyme within the same cell, each with separate biological functions (Otto and Smith, 1995). While previously noted, solid information on the forms of the enzymes responsible for PG biosynthesis in invertebrates is not yet available. Recently, the possibility of additional COX forms has arisen, surfaced by efforts to understand the pharmacology of acetaminophen (Tylenol in the USA and Paracetamol in the UK). While acetaminophen is an effective analgesic and antipyretic, it is a poor antiinflammatory drug and weak inhibitor of COX-1 and COX-2. The possibility of a COX-3 was first raised by Simmons and his colleagues

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Figure 3 Prostaglandin (PG) biosynthesis requires three enzymatic steps. First, AA (structure 1) is oxygenated to the unstable endoperoxide, PGG2 (structure 2). Second, a hydroperoxidase step reduces PGG2 to the more stable PGH2 (structure 3). Both of these enzymatic steps are catalyzed within a single protein, COX. PGH2 is converted into the classical prostaglandins, including PGE2 (4), thromboxane B2 (5), PGD2 (6), PGI2 (prostacyclin, structure 7), and PGF2a (8) by actions of cell-specific enzymes, as indicated. (Reproduced with permission from Stanley, D.W., 2000. Eicosanoids in Invertebrate Signal Transduction Systems. Princeton University Press, Princeton, NJ; ß Princeton University Press.)

(Simmons et al., 1999), who found that treating a murine macrophage line ( J774.2) with any of a variety of nonsteroidal antiinflammatory drugs induced expression of COX-2. The key finding was that, within the same cell line, the COX-2 induced by antiinflammatory drugs was more sensitive to inhibition with acetaminophen than COX-2 induced by LPSs. The authors postulated two enzyme activities with different properties. In another paper the authors suggested a COX-3, which is expressed as a variant of the COX-1 gene (Chandrasekharan et al., 2002). The story is further complicated by the suggestion that COX-3 is a variant of COX-2

(Botting, 2002). However, in all cases, the new COX expressed increased sensitivity to acetaminophen. This work opens the possibility of multiple isoforms of COX, all derived from just two separate genes (Warner and Mitchell, 2002) and also for the eventual discovery of rather specific therapeutics which act on specific tissues. Again, we have no information on COX isoforms in insects, although studies in this area may contribute important new knowledge on the enzymes responsible for PG biosynthesis. Figure 3 also shows the third enzymatic step in the biosynthesis of PGs. PGH2 is a substrate for several

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Figure 4 Two prostaglandins (PGs) are formed by nonenzymatic rearrangements of other PGs, rather than by enzymatic treatment of PGH2. PGA2 (3) is formed from PGE2 (1). D12-PGJ2 (4) is formed by a nonenzymatic dehydration of PGD2 (2). (Reproduced with permission from Stanley, D.W., 2000. Eicosanoids in Invertebrate Signal Transduction Systems. Princeton University Press, Princeton, NJ; ß Princeton University Press.)

enzymes responsible for converting PGH2 into the active PGs. The PGs D, E, and F are formed by three groups of enzymes, respectively, the PGD synthases, the PGE synthases, and the PGF synthases, (Urade et al., 1995). Two other PGs are formed directly from PGH2, thromboxane A2 and prostacyclin, also called PGI2 (Figure 3). Both of these compounds act in platelet aggregation, among other things. Thromboxase strongly induces platelet aggregation while PGI2 exerts the opposite action. PGI2 synthase is responsible for converting PGH2 into PGI2, and this enzyme is also a member of the cytochrome P450 superfamily (Tanabe and Ullrich, 1995). The structures of thromboxane A2 and PGI2 are a little different from the other PGs, and for this reason we see the term ‘‘prostanoid’’ used to describe all COX products. Two other PGs, D12-PGJ2 and PGA2 (Figure 4), are not formed directly from PGH2 (Negishi et al., 1995a). PGA2 is produced through a nonenzymatic rearrangement of PGE2. PGJ2 is a nonenzymatic dehydration product of PGD2, which is converted to the D12-PGJ2 in the presence of serum albumin. Most PGs, including PGE2, PGF2a, PGD2, PGI2, and thromboxane A2, express their actions through specific receptors located on cell surfaces. PGA2 and D12-PGJ2 operate through a different mechanism. These PGs are thought to be actively moved into cells via a transporter protein. An intracellular carrier molecule facilitates transport into the nucleus, where the PGs bind with thiol groups of nuclear proteins. The PG–protein complex then interacts with DNA, resulting in the expression of genes. Among their other biological actions, the A and J series PGs induce expression of genes for heatshock proteins in many normal and tumor cells.

4.9.2.2.3. The lipoxygenase (LOX) pathways Polymorphonuclear leukocytes are central to many mammalian inflammatory events. Samuelsson and his colleagues used these cells to investigate the possibility that AA could be metabolized into products other than PGs. They discovered that the main AA oxygenation pathways in leukocytes yielded 5-HETE and a series of products later called leukotrienes (Oates, 1982; Samuelsson, 1983). The mammalian AA LOXs produce a series of six products (Figure 5). These are 5-, 8-, 9-, 11-, 12-, and 15-HPETEs. Each of the LOXs is named according to the carbon that is oxygenated. Thus, 5-LOX yields 5-HPETE, and so forth (Pace-Asciak and Asotra, 1989). The LOX products are quickly taken into various metabolic systems, which yield other biologically active products. In one of the most common fates, the hydroperoxide products are quickly reduced to the corresponding HETE by various glutathione peroxidases, which are abundant in most mammalian cells. The HETEs are biologically active (Spector et al., 1988). For example, two products, 5- and 12-HETE, induce degranulation of human neutrophils. Similarly, 12-HETE is a potent chemoattractant for polymorphonuclear leukocytes. Several HETEs are active in various pathophysiological events, including proinflammatory processes. One of the LOX products, 5-HPETE, serves as substrate for biosynthesis of the LTs (Figure 6). The root LT is LTA4, an unstable epoxide of 5-HPETE. The enzyme responsible for this is called LTA4 synthase. The 5-LOXs from human leukocytes and mouse mast cells also express LTA4 synthase activity, and many workers believe both enzymatic steps are carried out by the same bifunctional enzyme.

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Figure 5 Lipoxygenase (LOX) is responsible for converting AA (1) into various hydroperoxyeicosatetraenoic acids (HPETEs) and hydroxyeicosatetraenoic acids (HETEs). Each of these compounds are formed by specific LOXs, identified by the positional specificity of the introduced oxygen. For example, 5-LOX yields 5-HPETE (2). The hydroperoxy acids may be reduced to their corresponding hydroxy acids by glutathione peroxidases. The remaining structures are identified by the position of the introduced oxygen: (3), 15-HPETE and 15-HETE; (4), 8-HPETE and 8-HETE; (5), 12-HPETE and 12-HETE; (6), 9-HPETE and 9-hydroxyeicosatetraenoic acid; (7), 11-HPETE and 11-HETE. (Reproduced with permission from Stanley, D.W., 2000. Eicosanoids in Invertebrate Signal Transduction Systems. Princeton University Press, Princeton, NJ; ß Princeton University Press.)

LTA4 is an unstable substance whose main function is to serve as a substrate for the biosynthesis of other LTs. There are two main pathways (Pace-Asciak and Asotra, 1989). The first is catalyzed by LTA4 hydrolase, which yields LTB4 (Figure 6). This LT is a proinflammatory mediator of host defense reactions in mammals. It activates polymorphonuclear leukocytes, myeloid cells, and mast cells. LTB4 also induces neutrophils to adhere to endothelial cell walls (Metters, 1995). Alternatively, LTA4 can be modified into the cysteinyl LTs or peptidoLTs (Figure 6). LTC4 synthetase catalyzes addition of glutathione, in covalent linkage, to C6, yielding LTC4. This LT can be converted to LTD4 by a single transpeptidase step which catalyzes hydrolysis of the terminal amino acid residue from LTC4. LTD4 undergoes another transformation to LTE4 by hydrolyzing the glycine residue from LTD4. This step is catalyzed by a dipeptidase. LTF4 can be formed by adding an amino acid residue. The cysteinyl LTs make up the slow-reacting substance of anaphylaxis (Samuelsson, 1983). The major biological action of these compounds is contraction of smooth muscles associated with respiratory and vascular systems, and with the alimentary canals of mammals. These actions are mediated through

specific receptors, of which two types are known (Metters, 1995). These are designated cys-LT1 and cys-LT2 receptors. Unlike the PG receptors, which exhibit marked specificity for each PG, the receptors for the cysteinyl LTs are much less fastidious. Both types of receptors have equal affinity for LTC4 and LTD4 and less affinity for LTE4. We have no information on receptors for LTF4. The LTs have not yet been considered in insect physiology. Beside the HPETEs, HETEs, and LTs, the LOX pathways can yield another suite of compounds, none of which is known from invertebrates. Similarly, products of the epoxygenase pathway mentioned in Figure 1 have not been discovered for invertebrates. These are considered in detail elsewhere (Stanley, 2000). Gerwick (1993) noted that the term eicosanoid is limited to oxygenated metabolites of a limited group of fatty acids, specifically C20 PUFAs. He suggested that a new term, ‘‘oxylipin,’’ was required to serve as a broader term for all oxygenated compounds formed from fatty acids of any chain length by reactions involving at least one step of a monooxygenaseor dioxygenase-dependent oxygenation. This broad word informs our appreciation of many fatty acidderived products in various animal systems. Another

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Figure 6 The biosynthesis and structures of leukotrienes. The pathways begin with conversion of AA to 5-HPETE by a 5-LOX. Leukotriene A4 synthase yields the unstable epoxide leukotriene A4 (LTA4). LTA4 may serve as substrate for leukotriene biosynthesis. In a one-step pathway, leukotriene hydrolyase yields leukotriene B4 (LTB4). The alternative pathway yields peptidoleukotrienes: leukotrienes C4 (LTC4), D4 (LTD4), and E4 (LDE4). (Reproduced with permission from Stanley, D.W., 2000. Eicosanoids in Invertebrate Signal Transduction Systems. Princeton University Press, Princeton, NJ; ß Princeton University Press.)

term, phytooxylipins, similarly describes a very wide array of oxygenated fatty acids that serve important roles in plant defense reactions (Blee, 1998). We may infer that the various forms of oxygenated fatty acids are crucial mediators in the life histories of most organisms. One area of most fundamental importance is reproduction, discussed in the next section.

4.9.3. Reproduction Appreciation of the biological significance of eicosanoids began with the discovery that a substance in unfractionated human semen causes contractions of uterine smooth muscle (Kurzrok and Lieb, 1930). Although we now know eicosanoids occur and exert biological actions in virtually every mammalian

tissue and body fluid, their influence on uterine muscle contraction, a function vital in reproductive biology, marks the discovery of the first known eicosanoid action. Release of egg-laying behavior in newly mated house crickets, Acheto domesticus, another action in reproductive biology, also marks discovery of the first known biological action of PGs in insects and other invertebrates (Destephano and Brady, 1977). Although only coincidental, it is quite intriguing because at a superficial glance, it appears that PGs mediate events in reproduction of mammals and a very distantly related invertebrate species. The occurrence and actions of PGs in insect reproductive systems have been reviewed (Stanley-Samuelson and Loher, 1986; Stanley-Samuelson, 1994a; Miller and Stanley, 1998a; Stanley 2000). These reviews

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remain useful because this is an area in which new information has been slow coming. This section presents an abbreviated synopsis of known information. The most detailed studies on the role of PGs in releasing egg-laying behavior come from the work of Werner Loher on the Australian field cricket, Teleogryllus commodus. Loher and his colleagues (Loher et al., 1981; Stanley-Samuelson et al., 1987) developed an ‘‘enzyme-transfer’’ model to account for transfer of a still-unidentified form of COX (along with its substrate, AA) from males to females via spermatophores. This model seems to hold for some, but certainly not all, cricket species. PGs may release egg-laying behavior in newly mated silkmoths, Bombyx mori, although some of the data on this species are not entirely convincing (Yamaja Setty and Ramaiah, 1979, 1980). PGs also may release egg-laying behavior in a few other insect species. Buprofezin is an insect growth regulator, which functions by inhibiting chitin biosynthesis and cuticle deposition in larvae of some insect species. Izawa et al. (1986) found that this compound inhibited egg-laying behavior in adults of the 28-spotted ladybird, Henosepilachna vigintioctopunctata. Similarly, Uchida et al. (1987) discovered that buprofezin inhibited egg-laying behavior and PG biosynthesis in adults of the brown rice planthopper, Nilaparvata lugens. The PG biosynthesis inhibiting action of the growth regulator suggested that PGs may release egg-laying behavior in these species. The occurrence among insect species of PGmediated egg-laying behaviors cannot be estimated in a reliable way. The model seems to apply to a few orthopterans, a lepidopteran, a coleopteran, and a hemipteran. Several points suggest that this is not a general model for insects. First, there are several insect species in which PG titers increase in female reproductive tracts after mating, but they do not release egg-laying behavior. Second, the role of PGs in releasing egg-laying behavior needs to be regarded in the context of individual mating systems (Thornhill and Alcock, 1983). In many species, eggs are formed, fertilized, and oviposited in separate time frames, and mating, per se, does not result in immediate egg-laying activities. Moreover, among those species in which mating does release egg-laying behavior, there are several mechanisms of releasing egg-laying behavior not involving PGs or other eicosanoids. Mechanical stimulation of egg-laying behavior is one example. PGs or PG biosynthetic activity are transferred from males to females of several insect species in which PGs do not influence egg-laying behavior (Stanley, 2000). Our question is: what is the meaning of this information? In general terms, one could

speculate that the PGs act in modulating and coordinating various details in reproductive physiology. The recent work by Medeiros and colleagues provides a specific example (Medeiros et al., 2002). As seen in other insect species, the ovaries of Rhodnius prolixus incorporate vitellogenin as well as several other specific proteins (see Chapter 3.9). Medeiros et al. (2002) tested the idea that eicosanoids influence the ovarian incorporation of Rhodnius heme-binding protein (RHBP). Ovaries of vitellogenic females were incubated in the presence of iodinated RHBP and either PGE2 or indomethacin, a COX inhibitor. Their results indicated that incubations in the presence of PGE2 downregulated RHBP uptake by up to 35%, while incubations in the presence of indomethacin upregulated RHBP uptake by up to 50%, both in comparison to control incubations. The authors also showed that the Rhodnius ovaries secreted PGE2 into the culture medium and that the amount of secreted PGE2 was reduced significantly in incubations conducted in the presence of indomethacin. Although the indomethacin did not influence the surface area nor the patency of follicle cells, indomethacin treatments did influence dephosphorylation of two ovarian proteins, an 18 kDa and a 25 kDa protein. Medeiros et al. (2002) concluded that eicosanoids influence the reproductive physiology of Rhodnius ovaries. Phosphorylation signal transduction pathways may be involved in PG modulation of RHBP uptake. The significance of this paper lies in its direct illumination of a previously unrecognized role of PGs in insect reproduction. Perhaps more important, however, the paper illustrates the point that many more-or-less subtle physiological steps in the overall process of producing progeny may be influenced by PGs and other eicosanoids. Aside from the roles of PGs in releasing egg-laying behavior in a few insect species and in modulating RHBP endocytosis in Rhodnius ovaries, eicosanoids act in various other aspects in reproduction of many invertebrates and lower invertebrates. This work on animals outside of Insecta has been treated elsewhere (Stanley-Samuelson, 1994a; Miller and Stanley, 1998; Stanley, 2000). It is worth noting that eicosanoids influence reproduction by acting at several levels of biological organization, as mentioned earlier for mammals. Actions such as modulating insect ovarian endocytosis of a protein (Medeiros et al., 2002) and many other eicosanoid actions in invertebrate reproduction (Stanley, 2000) are expressed at the cellular level, i.e., the eicosanoids influence cellular events. Other actions, including release of the egg-laying behavioral program are registered at the organismal level, even though they are probably driven by PG

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actions at the level of a few select neurons (Loher et al., 1981). Some eicosanoid actions take place at the population level. Sorensen et al. (1988), for example, reported that PGs of the F series act as the postovulatory pheromone in goldfish. There remains much to be discovered in the roles of eicosanoids in insect reproductive biology.

4.9.4. Eicosanoids in Ion Transport Physiology It is generally thought that cellular life evolved in aqueous environments. Cells live in more or less aqueous environments, and virtually all cells have evolved physiological mechanisms to maintain homeostasis of ion and water balance. Terrestrial insects tend to lose water to their environments, and they have mechanisms to restrict and offset water losses (Kirschner, 1991). Freshwater animals tend to maintain their extracellular fluid compartments hyperosmotic to their environments. Still other animals live in water subject to rapid changes in osmotic concentration, such as estuaries and rocky pools in intertidal zones. The osmotic homeostasis of all animals is subject to frequent challenges at the cellular and organismal levels. Many homeostatic mechanisms are expressed at the organismal level. The respiratory surfaces of all terrestrial animals, including insects, are internalized. The internalization of these surfaces is regarded as a major adaptation to terrestrial life. Terrestrial insects have a small layer of hydrocarbons and other lipids on their integument which helps reduce water loss. Some insects drink water to offset losses while other species can absorb water from humid atmospheres. Finally, some insects are able to sustain extreme dehydration. Larvae of the midge Polypedilum vanderplanki, which live in ephemeral pools on rocks in Africa, provide the best example of this. These pools evaporate in the dry season, and the larvae can tolerate nearly complete dehydration until the following rainy season (Hinton, 1960). Cellular mechanisms for maintaining water balance are integrated into organismallevel reactions to osmotic challenge. Vertebrate kidneys and the Malpighian tubules of insects and certain other invertebrates, for example, are influenced by diuretic and antidiuretic hormones which act to eliminate or conserve body water or solutes. To appreciate eicosanoid actions in ion transport, a brief overview of some mechanisms of solute transport is given (Alberts et al., 1994). The lipid bilayers of cellular membranes are fairly permeable to water and relatively impermeable to biologically important ions, such as chloride, sodium, potassium,

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magnesium, and calcium. These ions are transported across membranes by members of two classes of membrane-associated proteins, channels and carrier proteins. Channels are transmembrane proteins that form hydrophilic pores through membranes. The pores are generally selective, allowing specific ions, such as potassium or calcium, to pass through. Carrier proteins also bind specific solutes, which are transported due to conformational changes in the carriers. All channel-mediated and some carriermediated transport actions are driven solely by local electrochemical gradients known as passive transport or facilitated diffusion. Some carrier proteins are associated with an energy source, usually an ATPase. These carriers sometimes transport solutes against steep electrochemical gradients. Other carrier proteins act as coupled transporters in which a transport protein is responsible for movement of one solute with simultaneous movement of another. Symports move the two solutes in the same direction; antiports move the two solutes in opposite directions. For an example, the ubiquitous sodium– potassium antiport is responsible for actively pumping of sodium ions out of cells and potassium ions into cells. Passive and active transport mechanisms also are responsible for moving water across membranes. This brief glimpse allows us to regard the homeostasis of water and solute concentrations at the organismal and cellular levels as the outcome of regulated actions of specific intracellular, and often intramembrane, proteins. Eicosanoids are among the molecules which modulate homeostasis of water and solute concentrations. As usual, most of our knowledge of the roles of eicosanoids in the physiology of water and solute homeostasis comes from the studies on vertebrate systems, particularly various kidney and toad preparations. Nephrons are the operative cells in kidney water and solute transport, and eicosanoid biosynthesis and actions vary along the nephrons, as detailed by Bonvalet et al. (1987). Dalton (1977a, 1977b) provided the first recognition that eicosanoids act in the physiology of fluid secretion or ion transport in invertebrates. Fluid secretion by salivary glands isolated from the blow fly Calliphora erythrocephala can be stimulated by treating the glands with serotonin, a biogenic amine. In this system, the serotonin acts as an external hormone, and it is the natural ligand for salivary gland cell surface receptors. Serotonin-receptor interactions stimulate increased concentrations of intracellular cAMP in the salivary gland preparations (Berridge, 1970; Prince et al., 1972) and these increased intracellular cAMP concentrations lead to increased secretion of an isosmotic potassium-rich

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fluid. Dalton (1977a) investigated the influence of PGE1 on salivary gland physiology, showing that treating isolated salivary glands with low doses (10
7–10
9 M) of PGE1 did not alter their basal fluid secretion rates. Alternatively, these low doses of PGE1 attenuated the usual stimulatory influence of serotonin on fluid secretion rates. Dalton (1977b) suggested that PGE1 reduces intracellular cAMP concentrations by downregulating adenylate cyclase, with no influence on phosphodiesterase. It appeared that at least one eicosanoid, PGE1, plays an important physiological role in fluid secretion physiology in an invertebrate. In another early line of work with the locust rectum, Phillips and his colleagues suggested that eicosanoids may stimulate increased intracellular cAMP concentrations, which in turn lead to increased chloride transport (Phillips, 1980). Petzel and Stanley-Samuelson (1992) suggested that PGs modulate basal fluid secretion rates in Malpighian tubules of female yellow fever mosquitoes, Aedes aegypti. This hypothesis was tested in a series of fluid secretion assays based on the Ramsey protocol (Petzel, 1993). In each experiment Malpighian tubules were incubated in the presence of physiological buffer. After an equilibration period, half of the buffer was exchanged with the same volume of buffer containing an inhibitor of eicosanoid biosynthesis. First, the influence of eicosatetraynoic acid (ETYA), an AA analog with triple bonds in place of the usual double bonds, was assessed. ETYA inhibits many enzymes which process AA, including PLA2, LOX, and COX. In the presence of ETYA, basal fluid secretion decreased from about 1.0 nl min
1 in the pretreatment period to 0.55 nl min
1 in the experimental period. These findings supported the idea that PGs are involved in modulating basal fluid secretion rates in mosquito Malpighian tubules. Results of similar experiments with selective LOX and expoxygenase inhibitors indicated that products of these two pathways do not influence basal fluid secretion rates in Malpighian tubule preparations. Experiments with the COX inhibitor indomethacin, however, showed that PGs modulate basal fluid secretion rates. In the presence of 100 mM indomethacin, fluid secretion decreased from 0.9 to 0.5 nl min
1 in a dose-dependent manner. It was inferred from this work that PGs, but not LOX or epoxygenase products, are involved in maintaining basal fluid secretion rates in mosquito Malpighian tubules (Petzel and Stanley-Samuelson, 1992). The idea that PGs act in mosquito Malpighian tubules raised the issue of the occurrence and metabolism of AA in these tissues. Petzel et al. (1993) determined the presence of AA and PGE2 in mosquito

Malpighian tubules. They used immunohistochemistry to detect PGs (Howard et al., 1992; Petzel et al., 1993). The repeated dark-brown staining pattern showed the presence of PGE2 in principal, but not stellate, cells of the tubule. The principal cells are responsible for secreting fluid into the lumen of the tubule. A similar staining pattern was observed for PGE2 in Malpighian tubules of the yellow mealworm, Tenebrio molitor (Howard et al., 1992). PGF2a is also present in Malpighian tubules from the mosquito and the mealworm, shown by similar histological procedures. The PGF2a staining patterns are quite different from the patterns obtained for PGE2. For PGF2a, the staining is not restricted to the principal cells, but seems to be more or less evenly distributed among both major Malpighian tubule cell types. Hypothesizing that PGs act through G proteincoupled receptors, Petzel also conducted a preliminary investigation of the possibility that PGs stimulate increased cAMP biosynthesis in mosquito Malpighian tubules (D.H. Petzel, personal communication). The tubules were isolated from adult female mosquitoes, then incubated in mosquito saline. PGE2 was added to the saline and after selected incubation periods the tubules were frozen in liquid nitrogen and cAMP concentrations in the tubules were determined. This work showed that PGE2 stimulated a four- to fivefold increase in intracellular cAMP concentrations. Petzel also recorded the influence of PGE2 on fluid secretion rates. In these experiments, tubules were equilibrated as usual in mosquito saline, then PGE2 was added to the saline. PGE2 stimulated increased fluid secretion in mosquito Malpighian tubules, suggesting that PGs are among the regulatory elements in mosquito Malpighian tubule physiology. Van Kerkhove et al. (1995) conducted a similar line of work on Malpighian tubules from workers of the ant Formica polyctena. These findings are similar to the data with mosquito Malpighian tubules and support the notion that PGs act in insect Malpighian tubule physiology. Phillips (1980) implicated PGs in the physiology of the locust rectum. Radallah et al. (1995) continued this work using an everted sac preparation of the locust rectum to assess the influence of AA and PGE2 on water resorption. The everted sac is formed by tying one end of the rectum onto a catheter tube. Test compounds could then be injected into the sac, thereby exposing the hemolymph side of the preparation to the compounds. PGE2 stimulated increased fluid resorption in a dose-dependent manner. Maximal stimulation, 59% greater than controls, was obtained at about 10
9 M PGE2. They could also stimulate increased fluid resorption by adding AA, which resulted in a 79% increase in resorption at

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10
6 M AA. Similar experiments with aspirin and indomethacin also stimulated dose-dependent increases in fluid resorption. These data suggest that PGs and inhibitors of PG biosynthesis exert similar influences on the rectal preparations. Several possibilities help resolve this apparent paradox. For one, indomethacin restricts biosynthesis of all COX products, some of which exert inhibitory influences on cellular processes. The possible inhibitory PG may exert greater influence on overall fluid transport dynamics than the stimulatory PG. If so, inhibiting the biosynthesis of a possible inhibitory PG may create an apparent stimulation of fluid transport. The idea of considering possible inhibitory and stimulatory PG actions in the same system has not been explored. Radallah et al. (1995) determined the influence of AA and PGE2 on selected intracellular signal transduction messengers. To record the influence of AA, PGE2, and indomethacin on the influx of calcium into rectal cells they used a microfluorimetric technique. PGE2 and AA stimulated large increases in calcium influx, which decreased to baseline within minutes. As in the fluid transport experiments, indomethacin similarly provoked increased calcium influx. Pretreating the tissues with nifedipine, which blocks l-type calcium channels, completely blocked the effects of AA and PGE2 on calcium transport. This paper also reported that treating the locust rectum with AA or PGE2 stimulated substantial increases in phospholipase C (PLC) activity, which indirectly influences calcium transport (Berridge, 1993). Once again, aspirin and indomethacin similarly induced increased PLC activity. This apparently depends on the influx of calcium because an l-type calcium channel blocker strongly attenuated the influences of AA and PGE2 on PLC activity. They also showed that the PLC activity is associated with the release of inositol phosphates. Radallah et al. (1995) present a convincing postulate that AA and at least one eicosanoid, PGE2, are involved in modulating fluid transport in the locust rectum. The locust rectum is probably under primary regulation of the antidiuretic hormone, neuroparsin, which stimulates fluid resorption in the rectum. The hormone apparently acts through a G protein-coupled receptor, and it stimulates increased PLC activity. But it seems that the hormone also stimulates the release of AA from cellular PLs, which leads to increased PGE2. The PGE2 probably acts by an autocoidal mechanism to coordinate the cellular reactions to the central hormone. If so, PGE2 acts similarly in the insect rectum as in segments of the mammalian kidney (Bonvalet et al., 1987).

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PGs also modulate fluid secretion rates in salivary glands of the lone star tick, Amblyomma americanum (Qian et al., 1997). Tick salivary glands are osmoregulatory organs during host–parasite interactions. The salivary glands concentrate the nutrients associated with vertebrate blood and in the process they form a copious salt-rich saliva which is injected back into their hosts during feeding. The salivary glands are under nervous control and the neurotransmitter dopamine stimulates fluid secretion in isolated salivary glands (Sauer et al., 1995). Qian et al. (1997) investigated the influence of PGs on dopamine-stimulated fluid secretion. Compared to controls, treatments with the PLA2 inhibitor oleyloxyethyl phosphorylcholine (OOPC) and the COX inhibitor aspirin resulted in about 30–40% reductions in dopamine-stimulated fluid secretion rates. Longer incubations produced greater reductions and the influence of both inhibitors was expressed in a dose-dependent manner. The influence of OOPC was reversed by incubating the inhibitor-treated glands with 100 mM PGE2 or its stable analog 17-phenyl trinor PGE2. The PG treatments did not reverse the inhibitory influence of the COX inhibitors aspirin and diclofenac. Dopamine stimulates fluid secretion through G protein-coupled receptors resulting in increased intracellular cAMP concentrations (Sauer et al., 1995). Qian et al. (1997) determined the influence of PG biosynthesis inhibitors and of PGs on salivary gland cAMP concentrations. They found that the PLA2 inhibitor OOPC and the COX inhibitor indomethacin inhibited dopamine-stimulated increases in intracellular cAMP concentrations by about 25%. In the presence of OOPC, PGE2 and its analog stimulated 20–40% increases in the OOPC-treated cAMP concentrations. The eicosanoid system influenced neither fluid secretion nor intracellular cAMP concentrations in the absence of dopamine stimulation. The authors’ results provide strong support for their hypothesis that eicosanoids modulate dopamine-stimulated fluid secretion rates in tick salivary glands. Moreover, this work marks the first identification of physiologically functional PG receptors in an invertebrate system. Their data from binding studies support the view that the salivary gland receptor is functionally coupled to a stimulatory G protein. The authors concluded that the tick salivary gland expresses a functional PGE2 receptor that does not directly regulate adenylate cyclase. They speculated that the PGE2 receptor may influence calcium mobilization, which they confirmed in a subsequent paper (Qian et al., 1998). The biological significance of the salivary gland PGE2 receptor is linked

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to secretion, or exocytosis, of anticoagulant proteins, which are thought to facilitate blood-feeding. Qian et al. (1998) demonstrated this by incubating dispersed salivary gland tissue in the presence of PGE2, after which they recorded increased release of anticoagulant proteins. Taken together, work on blowfly salivary glands, locust rectum, insect Malpighian tubules, and tick salivary glands provide quite convincing evidence for the biological actions of eicosanoids in water and solute transport in representatives of arthropods. Eicosanoids similarly act in other invertebrates, lower vertebrates, and mammals (Stanley, 2000). It is suggested that regulating the actions of specific proteins involved in ion transport is a fundamental biological action of PGs and other eicosanoids in animals.

4.9.5. Insect Immunity 4.9.5.1. Introduction

The ability of organisms to defend themselves from parasites and pathogens is a fundamental aspect of biology. Many microbes produce and secrete enzymes and toxins that provide a measure of protection from other microbes. Plants elaborate sophisticated defense reactions to wounding as well as invasions by bacteria and fungi. Metazoan animals thrive in constant contact with pathogenic and nonpathogenic microbes, fungi, and parasites. An animal’s ability to defend itself from microbial invasions is known as immunity. Immunologists recognize separable categories of immune reactions, innate immunity and acquired (or adaptive) immunity. Innate immunity is made up of several protective mechanisms that function in a nonspecific way. In mammals and other higher vertebrates, clonal biosynthesis of antibodies to specific invading antigens is a main line of acquired, or adaptive, immunity. Invertebrates lack lymphocytes and immunoglobulins, and therefore do not produce specific antibody reactions to infections. On this basis invertebrates are often said, incorrectly, to lack immune systems. Rowley (1996) registered this incongruity, noting that some of the very important early experiments in immunology were conducted on invertebrates. Invertebrates and vertebrates express several forms of passive immunity, so named because they do not entail directed protective actions. These might include killing ingested organisms due to relatively harsh pH conditions (either strongly acid as in the mammalian stomach or very alkaline as in many insect midguts) in the alimentary canal. Foodborne

microbes may be killed by hydrolytic actions of digestive enzymes. The integument serves as a very effective barrier to invading microbes. Animals also express innate immunity to infections. Armstrong et al. (1996) recognized several expressions of innate immunity, some of which are seen in vertebrates and in invertebrates. Inducible antibacterial peptides of insects, defensins in insects and mammals, and the a2-macroglobulins, known in mammals and some arthropods, are elements of innate immunity. The innate immunity of insects is often divided into two major categories, cellular and humoral immunity. Cellular immunity involves direct interactions between circulating hemocytes and invading organisms (reviews: Brehelin, 1986; Dunn, 1986; Gupta, 1986, 1991; Boman and Hultmark, 1987; Lackie, 1988; Strand and Pech, 1995; Gillespie et al., 1997; Levine and Strand, 2002). Cellular defense reactions include phagocytosis, a form of endocytosis; nodulation, entrapping bacterial cells into clusters of hemocytes; and encapsulation of organisms too large for phagocytosis, such as eggs of parasitoid insects. Following capture, the internalized microorganisms are destroyed by intracellular killing mechanisms. Formation of oxygen radicals and nitric oxide are killing mechanisms within invertebrate cells (Conte and Ottaviani, 1995). These immune reactions create a general inflammatory response to injury, infection, and parasitization. Insect humoral immunity involves induced biosynthesis of antibacterial proteins (Tauszig et al., 2000; Hoffmann and Reichart, 2002). This very important area of inquiry has revealed a large number of antibacterial proteins and has generated new understanding of fundamental aspects of biology, including gene regulation. The chapter by Hultmark is the best entry point to literature on humoral immunity. Insects respond to large bacterial infections by nodule formation, the predominant response to bacterial infections (Dunn and Drake, 1983; Horohov and Dunn, 1983). Nodulation begins with entrapment of bacterial cells by granule-containing hemocytes. The granule-containing cells undergo degranulation, which releases proinflammatory chemicals. The nodulation process is completed by attaching layers of flattened phagocytes to the mature nodule. The last stage is a melanization process, leaving darkened, easily visible nodules attached to the inner sides of the body wall or various organs. Invading bacterial cells are topologically removed from circulation by effectively forming an impermeable wall between the bacterial mass and the remainder of the organism.

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Nodulation can be regarded as a form of cellular encapsulation, seen in response to infection by organisms larger than bacterial cells, such as parasites and other foreign materials. Wounds to the integument also evoke encapsulation reactions. As in nodulation, encapsulation begins with release of proinflammatory chemicals by degranulation of granule-containing cells. Again, the ending stages involve attaching layers of phagocytes, which flatten and melanize the capsule. 4.9.5.2. Eicosanoids in Insect Immunity

The inflammatory defense reactions just described have been studied quite extensively in many invertebrate species, particularly arthropods (Levine and Strand, 2002), although we have relatively little information on the biochemical molecules that mediate inflammation in invertebrates. Some mediators known from mammalian host defense reactions operate in invertebrates. Proinflammatory cytokines, or cytokine-like molecules, exist in invertebrates, including a bivalve mollusc, Mytilus edulis, a sea star, Asterias forbesi, and a urochordate, Styele clava, all reviewed by Rowley (1996). Downer and his colleagues showed that biogenic amines stimulate hemocytic inflammatory reactions in cockroaches (Baines et al., 1992; Baines and Downer, 1994) and waxmoths (Dunphy and Downer, 1994; Diehl-Jones et al., 1996). Similarly, it appears that eicosanoids, to which we now turn attention, also serve as crucial mediators of insect inflammatory reactions to bacterial infections. 4.9.5.2.1. Immune reactions to bacterial infection Eicosanoids exert many influences, in some cases stimulatory and in others inhibitory, on mammalian host defense systems. Based on this background, Stanley-Samuelson considered the possible actions of eicosanoids in insect immunity (Stanley-Samuelson et al., 1991). The results of these experiments, all using fifth instar Manduca sexta larvae, strongly supported the idea that eicosanoids mediate one or more cellular reactions to bacterial infections (Stanley-Samuelson et al., 1991). This paper prompted the question: which of the hemocytic defense reactions to bacterial infections are mediated by eicosanoids? Stanley and his colleagues hypothesized that eicosanoids mediate insect nodulation responses to bacterial infections (Miller et al., 1994). Nodulation can be assessed by counting numbers of melanized nodules within insect hemocoels following infection (described in detail by Miller and Stanley, 1998b). It was inferred from the results of these experiments that eicosanoids mediate one or more of the early steps in the

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nodulation reaction to bacterial infection (Miller et al., 1994). These two investigations with tobacco hornworms were the first experiments on eicosanoid actions in invertebrate immune systems. The assays for microaggregation and nodulation reactions are simple, and they facilitate investigation of a broader hypothesis. Specifically, do eicosanoids similarly act in nodulation reactions in other insect species? This question was tested in a series of similar exercises with other species. These include the tenebrionid beetle, Zophobas atratus (Miller et al., 1996), black cutworms, Agrotis ipsilon, and true armyworms, Pseudaletia unipuncta ( Jurenka et al., 1997), the silkworm, B. mori (Stanley-Samuelson et al., 1997), and caterpillars of the butterfly, Colias eurytheme (Stanley et al., 1999). The results of all the experiments with these species supported the hypothesis. Similar findings emerged from work with adults of the cricket Gryllus assimilis (Miller et al., 1999), adult cockroaches Periplaneta americana (Tunaz and Stanley, 1999) and adult 17-year periodical cicadas, Magicicada septendecim and M. cassini (Tunaz et al., 1999). Although these species do not establish an exhaustive representation of the Class Insecta, they make up a sufficient sampling of insects to suggest that PGs and other eicosanoids are key mediators of insect cellular immunity. Beyond the work described here, several other laboratories have investigated the influence of eicosanoids in insect immunity. Mandato et al. (1997) carried out a detailed investigation of eicosanoid actions in three discrete cellular processes within the overall nodulation reaction of the waxmoth larvae, Galleria mellonella. They found that eicosanoids mediate phagocytosis, cell spreading, and prophenyloxidase (PPO) activation in waxmoth larvae. Their work on PPO activation is quite interesting, because other groups found that eicosanoids do not influence PPO activation. Morishima et al. (1997) suggested a completely new role for eicosanoids in insect immunity. As mentioned earlier, the immune reactions of insects and other invertebrates to bacterial infections include humoral and cellular responses. The humoral responses include induced synthesis of antibacterial proteins, including cecropins and lysozymes. Cecropins are not found in the hemolymph in unchallenged insects, and the gene for this protein is somehow activated upon bacterial infection. Lysozyme occurs at low, constitutive levels in hemolymph, and expression of the lysozyme gene is upregulated following stimulation with bacterial cells or components of the bacterial cells. Morishima et al. (1997) suggested that eicosanoids mediate induction of the genes for cecropin

p0340

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and lysozyme in fat body of silkworm, B. mori. This work marks the recognition of a newly discovered eicosanoid action in invertebrate immunity, made all the more interesting in light of the recent discovery of a functional coupling between eicosanoid biosynthetic pathways and the immune deficiency (imd) pathway in Drosophila (Yajima et al., 2003). These authors reported that treatments with either of the two PLA2 inhibitors, dexamethasone or pbromophenacyl bromide, inhibited activation of the imd pathway. The inhibitory effects of these two inhibitors were attenuated by additional treatments with eicosanoid biosynthesis precursor fatty acids. Yajima et al. (2003) also found that AA alone did not activate the imd pathway, which indicated that eicosanoids participate in the activation of the imd pathway, but requires further LPS stimulation. 4.9.5.2.2. Immune reactions to pathogenic fungal infection All the work just mentioned is based on insect cellular defense reactions to bacterial infections. Dean et al. (2002) broadened the scope of this work with their hypothesis that eicosanoids mediate insect cellular reactions to the fungal pathogen Metarhizium anisopliae. They showed that treating tobacco hornworms with dexamethasone prior to challenge with fungal spores resulted in reduced number of nodules in reaction to the spores and that the influence of the drug could be reversed by treating experimental hornworms with AA. This was the first suggestion that eicosanoids mediate insect cellular reactions to fungal challenge. Lord et al. (2002) carried out a similar investigation using the fungal pathogen Beauveria bassiana. They reported that treating experimental tobacco hornworms with dexamethasone or the LOX inhibitors caffeic acid or esculetin or the COX inhibitor ibuprofen resulted in substantial reduction in number of nodules formed in reaction to fungal challenge. They also conducted a series of revealing rescue experiments. They found that the dexamethasone effect on nodulation could be reversed using the LOX product 5-HPETE, but not with the COX product PGH2. Moreover, the influence of caffeic acid and esculetin was reversed with 5-HPETE but the ibuprofen effect was not changed by treating experimental hornworms with PGH2. The authors inferred that products of the LOX pathways, but not the COX pathways, act in mediating hornworm immune reactions to B. bassiana. Lord et al. (2002) also noted that eicosanoids did not influence PPO activation, as seen in waxmoth larvae (Mandato et al., 1997). These two findings with different insect fungal pathogens add important new information to understanding signaling mechanisms in insect cellular immunity.

4.9.5.2.3. Immune reactions to parasitoid eggs Carton et al. (2002) added a new insight into insect immunity with their report that dexamethasone treatments inhibited cellular reactions to parasitoid invasion. In these experiments, larvae of the fruitfly Drosophila melanogaster were injected with dexamethasone, then exposed to adults of the parasitoid wasp Leptopilina boulardi. Compared to control larvae, in which approximately 90% of the parasitized larvae encapsulated the wasp eggs, about one-third of the larvae treated with 5 mg of dexamethasone encapsulated eggs and about 14% of larvae treated with 8 mg of dexamethasone encapsulated eggs. These results demonstrated a substantial dosedependent effect of dexamethasone on encapsulation of parasitoid eggs, from which the authors inferred that eicosanoids mediate cellular encapsulation reactions to parasitoid eggs. The information reviewed here may suggest that eicosanoids mediate cellular immune reactions to bacterial, fungal, and parasitoid challenge. It is suggested that the roles of eicosanoids in immunity may be another fundamental eicosanoid action in animals. More to the point of understanding insect immunity, however, it has been suggested that the immune system of D. melanogaster discriminates between bacterial and fungal challenge, seen with respect to differential activation of signal transduction pathways, which lead to expression of genes for antimicrobial peptides (Lemaitre et al., 1997). In light of this, it is noted that M. sexta cellular reactions to challenge by one fungal pathogen apparently depend on LOX, rather than COX, pathways, while reactions to bacterial challenge may be mediated by products of both pathways. As seen with the pathways for expression of genes for antimicrobial proteins, fungal and bacterial challenges seem to activate different pathways of eicosanoid biosynthesis. 4.9.5.2.4. Interactions between insect immune and neuroendocrine systems It was noted earlier that the signal transduction systems in insect cellular immunity include other important biomolecules, including peptides and biogenic amines. Beyond this, recent evidence points to interactions between the immune and endocrine systems in insect cellular immune reactions to infection. Wiesner et al. (1997) reported that apolipophorin-III stimulates immune reactions in G. mellonella. Halwani and Dunphy (1999) also noted that apolipophorin-III potentiates immune reactions in G. mellonella.Goldsworthy et al. (2003) considered another element of the neuroendocrine system in their study of adipokinetic hormone-I, apolipophorin-III, and eicosanoids in the locust,

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Locusta migratoria. They found that adipokinetic hormone and eicosanoids are important in signaling nodulation reactions while adipokinetic hormone and apolipophoren-III are crucial for stimulating PPO activation. In line with the report by Lord et al. (2002), eicosanoids do not seem to influence PPO activation in locusts. As noted elsewhere, there is room for many mediators in insect cellular immunity. Cellular reactions to immune challenge involve an unknown, albeit conceivably large, number of discrete cellular events (Miller et al., 1994, 1996; Stanley, 2000). Nodulation, in particular, undoubtedly involves many discrete cellular activities, only a few of which have been identified. While not meant to be an exhaustive list, these may include recognition of microbial cell wall components, producing and secreting various signal moieties which attract other hemocytes toward the site of infection, cell migration, various cell adhesion actions (microbe–hemocyte and hemocyte–hemocyte), cell spreading, and activation of prophenoloxidase. One of the major gains of investigating the roles of eicosanoids in cellular immunity is the possibility of recognizing additional cell actions and identifying which of many possible eicosanoids are responsible for signaling particular cell actions. 4.9.5.3. Testing the Eicosanoid Hypothesis

The idea that eicosanoids mediate cellular immune reactions to microbial and parasitoid challenge is a robust notion, which requires testing from a number of approaches. By and large, the evidence supporting the hypothesis has been circumstantial in nature. On one hand, we see that treating insects with pharmaceutical inhibitors of eicosanoid biosynthesis impairs cellular defense reactions to immune challenge. On the other hand, the ability of insect tissues, including immunity-conferring tissues, to produce PGs and other eicosanoids has been documented. However, this biochemical work does not, in itself, forge a link between eicosanoids and cellular immune signaling. In the following paragraphs, a few experiments designed to more directly test the eicosanoid hypothesis are reviewed. The outcomes of experiments with inhibitors of eicosanoid biosynthesis uniformly indicate that insects which had been treated with the inhibitors were impaired in their ability to clear bacterial cells from hemolymph circulation and in their ability to form microaggregates and nodules following infection. If the eicosanoid hypothesis makes sense, the impairments are due to the inability of hemocytes or other tissues to biosynthesize eicosanoids in response to microbial challenge. It should follow, then, that microbial infections stimulate production of various eicosanoids.

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Jurenka et al. (1999) investigated this possibility in a series of experiments with black cutworms. Cutworms were artificially infected by injecting heat-killed bacteria, Serratia marcescens, into their hemocoels. After 30 min incubations, hemolymph was collected and processed for eicosanoid extraction. The eicosanoids were derivatized with the fluorescent compound ADAM, then analyzed on high-performance liquid chromatography (HPLC) equipped with a flow-through fluorescence detector. This work revealed large increases in the titer of one PG, PGF2a, in hemolymph of experimental, but not vehicle-injected control insects. Moreover, pretreating experimental cutworms with selected COX inhibitors blocked the infection-stimulated increases in PGF2a. This work strongly supports the eicosanoid hypothesis because it indicates that insects increase PG production following bacterial challenge. Miller and Stanley (2001) considered the issue of signaling among immune tissues by investigating microaggregation reactions to bacterial challenge in isolated hemocyte populations. Hemocytes were prepared from tobacco hornworms, then challenged with lyophilized bacteria, S. marcescens. After selected incubation periods, generally 2 h, number of microaggregates were determined. These experiments documented the point that isolated hemocytes preparations are able to form microaggregates when challenged with bacteria. Results of similar experiments in which hemocytes were pretreated with various inhibitors of eicosanoid biosynthesis indicated that the microaggregation reactions also depend on eicosanoids. This work led to the suggestion that isolated hemocytes are competent to biosynthesize and secrete eicosanoids in reaction to bacterial challenge. This idea was confirmed by experiments with hemocyte culture media that had been conditioned by challenging the hemocytes with bacteria (Miller and Stanley, 2001). Hemocyte preparations were challenged with bacteria and the hemocyte media were filtered through microfilters to produce ‘‘conditioned medium.’’ Naive hemocyte preparations were then treated with the conditioned medium, and hemocyte microaggregation was determined. These experiments showed that exposing naive hemocyte preparations to conditioned medium was sufficient to generate microaggregation reactions in the naive cells. In another control experiment, it was observed that media, which was conditioned using hemocytes, that have been pretreated with dexamethasone or other inhibitors of eicosanoid biosynthesis, did not generate microaggregation reactions. However, this attractive view suggesting rapidly expanding discoveries of eicosanoid actions in

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invertebrate immunity is best viewed with skepticism (Stanley-Samuelson, 1994b). Several important issues should be addressed before the biological actions of eicosanoids in immunity are accepted. First, as mentioned elsewhere (Stanley-Samuelson and Dadd, 1983; Stanley-Samuelson et al., 1988; Stanley, 2000), the presence and significance of eicosanoid-precursor PUFAs is not well appreciated in insect biochemistry. Second, although there are several reports on the topic, there is very little information on eicosanoid biosynthesis in insects and other invertebrates. Third, the pharmacological fates of COX and LOX inhibitors in invertebrates is not known with certainty. All of these issues relate to various aspects of the biochemistry of eicosanoids, to which we now turn. 4.9.5.4. The Biochemistry of Eicosanoid Systems in Immune Tissues

The fat body and hemocytes are thought to be the immunity-conferring tissues of insects, and the eicosanoid systems in these tissues were of prime interest. Soon after the hypothesis that eicosanoids mediate clearance of invading bacterial cells from hemolymph circulation was tested, an investigation into the main points of eicosanoid biosynthesis in these two tissues from the tobacco hornworm, M. sexta was launched. 4.9.5.4.1. Polyunsaturated fatty acids The fatty acid compositions of total lipids and selected PL fractions prepared from hemocytes isolated from fifth instar tobacco hornworms were determined (Ogg et al., 1991). For these experiments, hemolymph was prepared by pericardial puncture, to prevent inadvertent activation of hemocytes. The hemocytes were pelleted by centrifugation, washed to eliminate hemolymph contamination, then processed for lipid analysis. As expected (Stanley-Samuelson et al., 1988), only traces (less than 0.1% of the fatty acids) of C20:3n
6, AA, or C20:5n
3 were associated with total hemocyte lipids, or with total hemocyte PLs. This finding opened the possibility that AA or other eicosanoid-precursor components are sequestered into particular PL fractions. Two major glycerophospholipids, phosphatidylcholine and phosphatidylethanolamine, were isolated. Analysis of the fatty acids associated with these PLs again revealed only traces of the eicosanoid-precursor components. With the idea that these low levels might be related to an environmental constraint, the artificial culture medium was analyzed, which revealed higher proportions of AA in the medium than in the hemocytes. Hence, it was concluded the presence of low proportions of AA in hornworm hemocytes is the usual biological condition (Ogg et al., 1991).

Certainly there are exceptions to this notion. One of the long-standing pillars of the field of animal lipids is that PUFAs are mostly associated with PLs, and less so with neutral lipids. This is generally true for insects, as well (Stanley-Samuelson and Dadd, 1983), but such ideas mut be tested. The triacylglycerol fractions prepared from tobacco hornworm heads yielded the highest proportions of AA ever recorded in a lepidopteran, i.e., AA constituted about 12% of the triacylglyerol fatty acids (Ogg and Stanley-Samuelson, 1992). A few other such pools of AA and C20:5n
3 have been recorded in insect studies, one of which certainly relates to PG biosynthesis. Stanley-Samuelson and Loher (1983) found AA in a special pool in spermatophores of the cricket T. commodus. The AA was present at about 24% of phosphatidylcholine fatty acids, and virtually absent from phosphatidylethanolamine fatty acids. The authors suggested that the AA was specially sequestered for transfer from male to female crickets during mating. Perhaps the high AA proportions in hornworm heads represents a pool of storage AA that can be drawn upon as needed. Obviously, this is a speculation which requires more concrete examination. More recently, Nor Aliza et al. (2001) reported the presence of high proportions of AA and C20:5n
3 in tissue PLs of adult fireflies, Photinus pyralis. They recorded AA at about 25% of PLs fatty acids in light organ and fat body from males, 13% for testes, and about 8% for midgut epithelia. Although such high proportions might indicate a very high potential for PG biosynthesis, preliminary experiments indicated that firefly tissues produced PGs at rates no different from other studied insect systems. Recording very low proportions of AA in M. sexta, fat body and hemocytes is consistent with the general picture of PUFAs in terrestrial insects. These low proportions elicit questions about insect fatty acid biochemistry. Do hemocytes actively maintain low proportions of AA and other eicosanoid-precursor PUFAs? This was investigated by tracing the incorporation and remodeling of radioactive fatty acids into hemocyte lipids (Gadelhak and Stanley-Samuelson, 1994). Four radioactive fatty acids, C18:1n
9, C18:2n
6, C20:4n
6, and C20:5n
3, were used in separate incorporation experiments. Consistent with the general background of animal lipid biochemistry, the Manduca hemocytes incorporated all four of the radioactive fatty acids into cellular complex lipids. Of the four, C18:2n
6 was most efficiently incorporated into PLs, while C18:1n
9 and C18:2n
6 were efficiently incorporated into triacylglycerols. About 1–3% of the starting radioactivity in C18:1n
9 was recovered

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in diacylglycerols and monoacylglycerols. Very little of the other fatty acids was incorporated into these two fractions. The incorporation patterns generally agreed with the fatty acid composition of hemocytes. The incorporated fatty acids were redistributed among lipid fractions in hornworm hemocytes during longer incubation periods (Gadelhak and Stanley-Samuelson, 1994). This redistribution is due to selected hydrolysis of some components from PLs, allowing incorporation of other components. Chilton and Murphy (1986) documented this remodeling process in human neutrophils, showing that after initial incorporation into ester-linked PLs, radioactive AA was selectively remodeled into etherand plasmalogen-linked PL pools over time. These PL fractions were not considered in our studies because they have not yet been sufficiently detailed in insect systems. Remodeling of incorporated fatty acids from PLs to triacylglyerols was recorded (Gadelhak and Stanley-Samuelson, 1994). After longer incubations, the radioactivity recovered in the PL fraction declined, with concomitant increases in radioactivity in the triacylglycerols. For AA, the radioactivity recovered in PLs declined to about 97% at 20 min, and to about 83% by 120 min. This directional shift was not seen with incorporated C18:2n
6 and C18:1n
9, although some of the radioactivity associated with these fatty acids was shifted between these two major cellular lipid fractions. A similar picture emerged from the analysis of selected PL fractions. After 5 min incubation periods, most of the radioactivity associated with AA was recovered in phosphatidylcholine. With longer incubations, the proportions of radioactivity recovered in phosphatidylcholine declined, with attending increased radioactivity in phosphatidylethanolamine. After 120 min incubations, slightly more radioactivity associated with AA was detected in phosphatidylethanolamine, rather than phosphatidylcholine. Again, these results support the idea that PUFAs are selectively remodeled among the complex lipid fractions in hornworm hemocytes. Similar analyses of other insect tissues have not yet been carried out, but nonetheless, remodeling dynamics of this nature are to be expected. Again, these processes help understand the low proportions of AA and other eicosanoid-precursor PUFAs in terrestrial insects. AA is present in fat body from larvae of the beetle Z. atratus (Howard and Stanley-Samuelson, 1996; Miller et al., 1996). Similarly, AA is present in trace levels in black cutworms, A. ipsilon, true armyworms, P. unipuncta ( Jurenka et al., 1997), in silkworms, B. mori (Stanley-Samuelson et al., 1997), and in

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adults of the cricket G. assimilis (Miller et al., 1999). Relative to the work on waxmoth hemocytes (Mandato et al., 1997), AA and C20:5n
3 were confirmed in these insects during early studies of fatty acid nutritional metabolism (Stanley-Samuelson and Dadd, 1984; Stanley-Samuelson et al., 1988). Hence, one element required for eicosanoid biosynthesis in invertebrate immune tissues is present, albeit at low levels, in all systems in which eicosanoids are present. 4.9.5.4.2. Phospholipase A2 is the first step in eicosanoid biosynthesis Two of the eicosanoid biosynthesis inhibitors used in the immune studies, namely dexamethasone and p-bromophenacylbromide, are thought to act at the level of PLA2. These two reagents do not directly inhibit eicosanoidbiosynthetic enzymes, but inhibit the overall process of eicosanoid biosynthesis by arresting the release of AA or other PUFA from cellular PLs. In the absence of free substrate, eicosanoid biosynthesis cannot proceed. Because these two inhibitors attenuate cellular and humoral immune reactions to bacterial infections and the influence of one of these, dexamethasone, can be reversed by treating experimental insects with free substrate, AA, it was proposed that an intervening PLA2 step must be operative in the eicosanoid-mediated immune reactions to bacterial infections. The PLA2s are responsible for hydrolyzing the acyl moiety from the sn-2 position of glycerophospholipids and an intracellular PLA2 in Manduca fat body has been characterized (Uscian and StanleySamuelson, 1993). In brief, phosphatidylcholine with radioactive AA esterified at the sn-2 position was prepared in the form of substrate vesicles to facilitate enzyme activity. Manduca fat body homogenates were centrifuged to produce microsomalenriched fractions used as enzyme sources. The enzyme was incubated with substrate vesicles, then total lipids were extracted from the reaction mixtures. The lipid extracts were separated and fractions associated with unprocessed substrate, free fatty acids, and diacylglycerol were radioassayed. PLA2 activity was calculated from the amount of radioactivity associated with the free fatty acid fraction. The fat body PLA2 was sensitive to the usual biophysical parameters and the calcium requirements for this enzyme were assessed. For this experiment, the fat body was homogenized and centrifuged in calcium-free buffer containing the calcium chelator ethylene glycol bis (B-aminoethyl/ether)-N,N,n0 , N0 -tetraacetic acid (EGTA). Reactions run in the presence or absence of calcium yielded similar results, from which we inferred that the Manduca

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fat body PLA2 is a calcium-independent enzyme, or possibly one with very low stringency calcium requirements. To test this hypothesis, enzyme preparations were dialyzed against 200 volumes of EGTA buffer and enzyme activity was assessed. Again, rigorous removal of calcium did not reduce enzyme activity. A similar characterization of a PLA2 in hemocytes from fifth instar larvae was reported by Schleusener and Stanley-Samuelson (1996). For these experiments, hemolymph was collected by pericardial puncture, diluted in buffer, the hemocytes rinsed in fresh buffer, and the cells were homogenized by sonication. Microsomal-enriched fractions were prepared from the sonicates and these were used as the enzyme source. Again, the hemocyte PLA2 was sensitive to substrate concentration, protein concentration, and pH. As seen in virtually all mammalian preparations, the hemocyte enzyme was inhibited in the presence of 5–50 mM OOPC. The calcium requirements of the hemocyte PLA2 were determined, and as in the fat body preparations, all hemocyte preparation steps were carried out in buffer containing EGTA. Enzyme activity increased with increasing concentrations of EGTA, which suggested that the chelator removes from the reaction medium an ion that downregulates the hemocyte PLA2 (Schleusener and Stanley-Samuelson, 1996). Based on findings with some mammalian cPLA2s, the idea that the hemocyte PLA2 exhibits a selectivity for arachidonyl-associated PLs was considered. In these experiments, enzyme sources were prepared as usual, then incubated with either of the two substrates, one a palmitoyl-associated phosphatidylcholine and the other an arachidonyl-linked phosphatidylcholine. Enzyme activity with the palmitoyl-associated substrate was reduced by about 40% relative to activity with the arachidonylassociated substrate. It is more appropriate to assess substrate specificity using purified, or at least highly enriched, enzyme preparations since most animal cells have several forms of cytosolic PLA2, and experiments meant to detect the substrate preferences of a subset of these enzymes can yield ambiguous results. In this case, however, the 40% reduction in enzyme activity indicates that at least one PLA2 in Manduca hemocytes has a fairly strong preference for arachidonyl-linked PL substrate. This enzyme may regulate eicosanoid biosynthesis in hemocytes. These preliminary investigations serve to document both the presence of a cytosolic PLA2 in hornworm fat body and the fact that hemocytes can hydrolyze AA from the sn-2 position of cellular PLs. These results are preliminary because

documenting the presence of an enzyme in fat body and hemocytes does not demonstrate unequivocally that the enzyme is an integral part of the eicosanoid-mediated cellular reactions to bacterial infections, although recent work supports the idea that PLA2 is a key step in eicosanoid biosynthesis following bacterial infection. The circumstantial evidence on the influence of PLA2 inhibitors on cellular and humoral immune reactions has been bolstered with more direct experiments. Given the very low levels of AA in Manduca immune tissues, some of these experiments could be better designed using other invertebrate systems. For now, however, it can be said that the tobacco hornworm expresses cPLA2s, and some forms of these enzymes may be involved in immune reactions to bacterial infections. 4.9.5.4.3. Eicosanoid biosynthesis in immune tissues While eicosanoid biosynthesis has been demonstrated in a number of invertebrate systems, the idea that eicosanoids mediate cellular and humoral immune reactions to bacterial infections in insects opens a window of opportunity for investigation. Eicosanoid biosynthesis in Manduca fat body and hemocyte preparations has been recorded, but before considering this work, an important issue should be considered. The methodology of tracing PG biosynthesis in invertebrates has not yet matured to a series of standard techniques. Although most methods in essence amount to reacting an enzyme source with radioactive AA, there are a few subtle points which deserve attention because they can influence the outcome of such experiments. In work with the fat body (Stanley-Samuelson and Ogg, 1994), the eicosanoid biosynthesis reaction mixtures included radioactive AA emulsified in buffer containing a cofactor cocktail (2.4 mM reduced glutathione, 0.25 mM hydroquinone, and 25 mg hemoglobin in each reaction). The reactions were preceded by 3 min preincubations with all reaction components except the enzyme sources. The reactions were started by adding the microsomal-enriched enzyme source, and stopped by acidification and extraction of products. The radioactive PGs were separated by thin-layer chromatography (TLC) (Hurst et al., 1987). Chromatography with this system differs from usual procedures because the plates are placed into the solvent immediately after the solvent is transferred to the developing chamber. There is no equilibration period in this system, which requires about 2.5 h to develop. In some experiments, the reaction products were also separated by two-dimensional TLC. Bands corresponding to authentic standards were transferred to vials, and the radioactivity in each fraction

Eicosanoids

was determined. PG biosynthesis was calculated from the amount of radioactivity in each fraction. PUFAs are fairly unstable molecules in oxygen atmospheres and experimental artifacts can result from spontaneous oxygenation reactions, or autoxidation, during reactions and subsequent work up steps. Autoxidation was determined by conducting ‘‘zero-time’’ reactions. In these control experiments, the substrate was preincubated in parallel with the experimental tubes. At time zero, when the enzyme source would ordinarily be added, the control preincubations were stopped and extraction and chromatographic separation followed. Zero-time reactions were done in every experiment, and values from these control experiments were used to correct for autoxidation in the biosynthesis reactions. The fat body preparations yielded four PGs: PGF2a, PGE2, PGD2, and PGA2. Under most experimental conditions, PGA2 was the predominant fat body product. The enzyme preparation was sensitive to routine biophysical parameters although the literature on PG biosynthesis in invertebrates presents an ambiguous picture with respect to optimal reaction times. Drawing on the mammalian data (Smith et al., 1996), the first step in PG biosynthesis is catalyzed by COX, which undergoes ‘‘suicide’’ inactivation after about 1400 catalytic operations. The suicide inactivation is regarded as an autoregulatory mechanism, which imposes an upper limit on cellular potential to biosynthesize PGs. PG biosynthesis therefore occurs in short, very rapid bursts of enzyme activity. However, the overall dynamics of maintaining appropriate PG titers within cells includes PG biosynthesis and degradation. This could produce two phases of PG biosynthesis in in vitro reactions and possibly in intact cells. Early in the reactions, the rapid bursts of biosynthesis would exceed PG degradation, thereby favoring product accumulation. During the second phase of the reaction, the suicide inactivation of COX would lead to decreased PG biosynthesis and product degradation would exceed product formation. The overall result of this asymmetry in the reaction progress would be registered as higher product accumulation in shorter reaction periods which decreases during longer reaction periods. In work with the Manduca fat body, an initial burst of PG biosynthesis which peaked at about 1 min was observed. Thereafter, product accumulation decreased over the next 9 min. These findings agree with mammalian data, and also with the time course of PG biosynthesis seen in housefly preparations. Wakayama et al. (1986) also observed rapid PG biosynthesis during 2 min incubations. After the first 2 min, there was a very gradual increase in

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product formation over the following 58 min. In contrast, Brenner and Bernasconi (1989) reported a linear increase in PGE2 biosynthesis over a 60 min time course. In their characterization of PG biosynthesis by spermatophore content from males of the Australian field cricket, T. commodus, Tobe and Loher (1983) also found a linear increase in synthesis over 60 min reaction periods. The systems in which longer reaction times promote increased product formation may point to important differences between the well-established mammalian studies and the emerging information on invertebrate eicosanoid systems. All of the biological investigations into the roles of eicosanoids in cellular and humoral immunity described in this section were based on the use of pharmacological inhibitors of eicosanoid biosynthesis. Some of these inhibitors designed for studies on mammalian tissues reduce PG biosynthesis in hornworm tissues. In these experiments, the fat body preparations were incubated in the presence of selected doses of the pharmacological reagents, and the reaction products were then extracted and analyzed. The fat body preparations were very sensitive to the COX inhibitors, indomethacin and naproxin. At the low dosage of 0.1 mM, indomethacin and naproxin reduced total PG biosynthesis by about 80%. Higher dosages virtually abolished PG biosynthesis. Along with inhibition of PG biosynthesis, reactions with naproxin yielded substantial levels of a radioactive product, tentatively identified as the LOX product 15-HETE. The influence of naproxin on increased LOX activity occurred in a dose-dependent manner. In the presence of increasing naproxin concentrations, the decreasing COX activity was accompanied by increasing LOX activity. While indomethacin effectively inhibited COX activity, there was no accompanying increase in LOX activity. It may be presumed that the indomethacin also inhibits the insect LOX, as it does in some mammalian preparations. Tobacco hornworm hemocyte preparations also are capable of eicosanoid biosynthesis (Gadelhak et al., 1995). Pools of hemocytes were washed, then homogenized by sonification, microsomal-enriched preparations were prepared by centrifugation, and eicosanoid biosynthesis assayed. The hemocyte preparations yielded two major products, the COX product PGA2 and the LOX product 15-HETE. The identification of 15-HETE is based on a single TLC step, and undoubtedly that fraction contains more than one LOX product. For this reason, this product is simply noted as total LOX activity. Two inhibitors of eicosanoid biosynthesis influenced eicosanoid biosynthesis by the hemocyte

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preparations. The COX inhibitor naproxin reduced COX activity by 88% at 0.1 mM and by 96% at 1.0 mM. Contrary to results with the fat body, the hemocyte LOX activity was not enhanced in the presence of naproxin. Reactions carried out in the presence of the LOX inhibitor esculetin also yielded reduced COX and LOX product formation. At 0.1 mM, esculetin nearly inhibited all eicosanoid biosynthesis. These studies support the view that eicosanoid biosynthesis observed on TLC is not an artifact of substrate autoxidation. However, the influence of these eicosanoid-biosynthesis inhibitors is quite instructive. While these are well-characterized pharmacological agents in mammalian systems, their actions in invertebrate systems may differ in important ways. For example, indomethacin, a COX inhibitor, potently inhibited cellular defense reactions and COX activity in Manduca fat body preparations. This compound did not inhibit PG biosynthesis in preparations of male reproductive tracts from the cricket A. domesticus (Destephano et al., 1976). More to the point, naproxin is accepted as a specific COX inhibitor. Results with the hornworm fat body preparation would suggest that this is so in invertebrates as well, because low doses of naproxin (0.1 mM) effectively inhibited COX, but not LOX, activity. These results suggest that the influence of eicosanoid-biosynthesis inhibitors should be investigated in each species, and each tissue within a species, to ensure that the inhibitors act as they do in vertebrate experiments. This comment is quite relevant to work on immunity. On the basis of a single experiment with naproxin, one might conclude that COX activity is essential to cellular immune reactions to bacterial challenge. However, the results of our biochemical experiments with hemocyte preparations make it clear that naproxin may act by inhibiting LOX as well as COX activities. This is one of the reasons for conducting experiments with several separate inhibitors. Eicosanoid biosynthesizing enzymes are rather uniformly distributed within mammalian cellular fractions, with COXs exclusively associated with endomembrane fractions of cells. The subcellular localization of COX and LOX activities in hemocytes can be assessed by determining eicosanoid biosynthesis in the mitochondrial, microsomal, and cytosolic fractions. The COX activity was unevenly distributed among the three cellular fractions, about 5% in the mitochondrial fraction, 58% in the microsomal fraction, and about 36% in the cytosolic fraction. Most of the LOX activity (87%) was recovered in the cytosolic fraction, and the remaining 13% in the microsomal fraction. These data indicate

that insect COXs do not have the same subcellular distribution as one finds in mammals. Although the broad themes are similar, there is substantial variation in eicosanoid systems among mammalian species and among tissues within a mammalian species. Relative to invertebrates, much more variation will emerge as research activities which continue to generate new information. Therefore, it is important to investigate at least some aspects of eicosanoid biosynthesis in several insect species in which eicosanoids play a role. AA and other C20 PUFAs as well as PG biosynthesis capacity is present in tissues from the tenebrionid beetle Z. atratus (Miller et al., 1996). Similarly, AA is present in fat body PLs from true armyworms and black cutworms (Jurenka et al., 1997). The fat body from these larvae also expressed an intracellular PLA2 that can hydrolyze AA from cellular PLs. Fat body preparations from both species are able to convert radioactive AA into PGA2, PGD2, PGE2, and PGF2a. A similar line of documentation experiments showed the presence of eicosanoid-biosynthesizing enzymes in fat body from silkworms, B. mori (Stanley-Samuelson et al., 1997). Miller et al. (1999) recorded traces of AA in the fat body of adult crickets, G. assimilis, as well as biosynthesis of three eicosanoids by fat body preparations, PGA2, PGE2, and a hydroxyeicosatetraenoic acid. As in the silkworm fat body preparations, the LOX product(s) was the major product produced by the cricket fat body preparations. The point of these exercises is to document the presence of an eicosanoid-biosynthesizing system in insects thought to use eicosanoids in cellular immune signal transduction mechanisms. Even at this superficial level of analysis, however, substantial differences are evident in the eicosanoid systems among these few insect species. More detailed characterizations of the elements of eicosanoid biosynthesis in these species will undoubtedly reveal more differences. The biochemistry of these systems merits considerably more research attention. 4.9.5.4.4. The pharmacology of eicosanoid biosynthesis inhibitors There is very little knowledge concerning the movements or metabolic fates of eicosanoid biosynthesis inhibitors in invertebrates. Murtaugh and Denlinger (1982) maintained a group of house crickets on diets with indomethacin. After several days on these diets, they found reduced levels of PGE2 and PGF2a in testes from the experimental males and spermathecae from mated females. These findings suggest that indomethacin can move from the alimentary canal to at least two tissues in house crickets and also suggest the possibility of substantial biochemical differences between

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the action of these compounds in whole animals and enzyme preparations. These issues motivated the study of pharmacology of indomethacin in Manduca. Using fifth instar larvae, radioactive indomethacin was injected into the hemocoels of the animals and its movement, excretion, and metabolism examined. Over 99% of the radioactivity associated with injected indomethacin was cleared from the hemolymph circulation within the first 3 min after injection. The indomethacin was rapidly taken up by hornworm tissues. Radioactive indomethacin was recovered from all tissues analyzed, including integument, ventral nerve cord, salivary gland, fat body, Malpighian tubules, and gut epithelium. The greatest amount of radioactivity was recovered from fat body, the largest tissue. However, when normalized to wet tissue weight, most radioactivity was recovered from the salivary glands. These results indicate that indomethacin is probably distributed among all tissues in this insect, but not uniformly, as seen in mammals. Less radioactivity was recovered from integument and nerve cord, while more was recovered from fat body, silk gland, and Malpighian tubules. This is also consistent with the pharmacology of indomethacin in mammals. The extraction system used to recover indomethacin and its metabolites from hornworm tissues provided information on the metabolic fate of indomethacin. Indomethacin is highly soluble in chloroform and other organic solvents, and is virtually insoluble in water. Rigorous multiple extraction procedures yielded about 90% of the recovered radioactivity in the collected organic phases. The remaining 10% of the radioactivity was detected in the aqueous phases. This 10% represents polar metabolites of indomethacin, taken to indicate that about 10% of the injected indomethacin was metabolized to water-soluble products. There was no evidence for the metabolism of indomethacin in most tissues, including integument, nerve cord, fat body, Malpighian tubules, and gut epithelium. Virtually all of the radioactivity recovered from these tissues cochromatographed with authentic indomethacin on TLC. Contrary to results with other tissues, the salivary gland produced at least one polar product of indomethacin, which made up no more than 25% of the starting material. Hence, the material taken up into virtually all hornworm tissues is present as indomethacin. The excretion of radioactive indomethacin was recorded by collecting frass. In this experiment, radioactive indomethacin was injected into the hemocoel of fifth instar larvae. The insects were held in individual cups, and frass pellets were collected every 2 h for

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the following 40 h. About 56% of the injected material was recovered over the 40 h incubation period. A trace of radioactivity appeared in the frass as early as 2 h after injection and almost half of the injected radioactivity was recovered between 4 and 12 h. These data reveal that substantial amounts of injected indomethacin remain in the hornworm tissues in its original form during the first 12 h after treatments. For purposes of experimental design, the inhibitor was present in the hornworm throughout the time course of our immunology experiments. The products extracted from the frass were analyzed and three to five indomethacin metabolites were present in frass, accounting for about 30% of the recovered radioactivity. Since enteric microbes may have been responsible for the apparent metabolism, radioactive indomethacin was incubated with freshly collected frass pellets and the data showed that about 10% of the starting material was metabolized into more polar products after 40 h. This result suggests that one or more factors in hornworm frass are responsible for indomethacin metabolism prior to excretion. The composite data indicate that indomethacin is an appropriate probe for assessing the roles of eicosanoid biosynthetic pathways in the cellular immunity of tobacco hornworms, rather than insects or invertebrates, because there are substantial differences in the pharmacology of indomethacin among mammalian species. For example, dogs and guinea pigs require about 20 min to clear 50% of injected indomethacin doses from the circulation (Yesair et al., 1970), whereas rats require about 4 h (Hucker et al., 1966). The distribution of indomethacin among tissues within mammals also differs among species. Rats maintain higher concentrations of indomethacin in blood circulation than in tissues at all times after injection. In guinea pigs, indomethacin is concentrated from blood into liver, kidney, and small intestine. The excretion of indomethacin also differs among mammals (Hucker et al., 1966; Yesair et al., 1970). Nearly all injected indomethacin is excreted in feces in dogs, while rabbits excrete most injected indomethacin in the urine. Guinea pigs and humans excrete about half of the injected indomethacin in urine, the remainder in feces. These findings emphasize differences in distribution, metabolism, and excretion of injected indomethacin among mammals. Similar differences are to be expected among invertebrates. Indeed, the different biochemical effects of indomethacin on eicosanoid biosynthesis in insect systems have been discussed. Overall, this work points to the importance of understanding the pharmacology of eicosanoid biosynthesis inhibitors in the work on assessing the roles of eicosanoids

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in invertebrates, and the pharmacology of inhibitors in general.

4.9.6. Emerging Topics During the time Dadd (1985) was developing his review of insect nutrition, appreciation of the biological significance of PGs and other eicosanoids in insects or other invertebrates was an emerging topic in insect biochemistry and physiology. There was virtually no background literature on the presence or meaning of AA in insect tissue lipids (Stanley-Samuelson and Dadd, 1983) and very little idea of PG action in insects. Understanding of eicosanoids in insect biology has grown in the last two decades, mostly due to studies that revealed new – or emerging – information. In this section several emerging areas have been highlighted. 4.9.6.1. An Insect Pathogen Inhibits Eicosanoid-Mediated Immunity

The significance of the hypothesis that eicosanoids mediate insect cellular immunity is the identification of a previously unrecognized and crucial signal transduction system in insect immunity. The biological actions of eicosanoids involve a suite of enzymes responsible for eicosanoid biosynthesis and, presumably, specific receptors and intracellular receptor interactions with effector systems. These enzymes, receptors, and intracellular proteins all represent potential novel targets that may ultimately be exploited in the ongoing development of new insecticides. Given the diversity of microbial organisms, it is not surprising to see that at least one insect pathogen has already exploited the inhibition of eicosanoid biosynthesis as a mechanism to downregulate insect immunity. The nematode Steinernema carpocapsae lives in a symbiotic relationship with the bacterium Xenorhabdus nematophilus from which both partners gain advantage. The nematode serves as a productive environment for growth of the bacterium and X. nematophilus serves the nematode by killing newly invaded insect hosts. The biology of this and other bacterium–nematode–insect systems has been well documented (Kaya and Gaugler, 1993). Park and Kim (2000) recently illuminated a new aspect of these systems with their report that eicosanoids attenuate the lethality of X. nematophilus in the insect Spodoptera exigua. The authors inferred that the bacterium somehow inhibits eicosanoid biosynthesis in newly infected host insects, thereby impairing the insect cellular immune reactions to the presence of the bacterium. It was hypothesized that the bacterium inhibited eicosanoid biosynthesis in insect immune tissues by

restraining the action of PLA2, the first step in eicosanoid biosynthesis. Testing this hypothesis began by showing that bacterial infections stimulate increased PLA2 activity in hemocytes from tobacco hornworms (Tunaz et al., 2003). The significance of this work is that it supports the view that bacterial infections lead to increased eicosanoid biosynthesis (as shown by Jurenka et al. (1999)) by stimulating hemocytic PLA2 activity. It was subsequently found that, as Park and Kim (2000) originally suggested for S. exigua, X. nematophilus impairs nodulation reactions to bacterial infection in tobacco hornworms by inhibiting eicosanoid biosynthesis (Park et al., 2003). This work showed that hornworms challenged with living bacteria produced far fewer nodules than did hornworms challenged with heatkilled bacteria. The immunity-impairing influence of challenge with living bacteria could be off-set by treating experimental hornworms with AA. It may be inferred that X. nematophilus inhibits eicosanoid biosynthesis in all of its host insect species. It appears from this work (Park et al., 2003) that the bacterium X. nematophilus secretes a specific product that inhibits eicosanoid biosynthesis. To investigate this, X. nematophilus cells were cultured, then separated from their culture medium by centrifugation; the medium was then fractionated into an aqueous and an organic fraction. After concentrating the extracts, each fraction was tested for its influence on nodulation. The organic fraction inhibited nodulation reactions to living bacteria while the aqueous fraction had no influence on nodulation. The organic fraction was fractionated into five discrete fractions by column chromatography. After testing each of the five, nodulation-inhibiting activity was present in only one. Moreover, results indicate X. nematophilus cells and its immunity-impairing product specifically inhibits PLA2 in tobacco hornworm hemocytes (Y. Park et al., unpublished data). The identity of the inhibitory factor should be determined in the near future. This work revealed important new information on the biochemical mechanisms, which underlie the relationships among certain entomopathogenic nematodes, their symbiotic bacteria, and their insect hosts. It also strongly supported the hypothesis that eicosanoids mediate insect cellular defense reactions to microbial infection. Another aspect of the significance of these findings is that they indicate the potentials for discovery of mediating mechanisms in insect physiology and biochemistry. 4.9.6.2. Bioactive Oxylipids from Linoleic Acid

In addition to oxygenation of AA via the COX or LOX pathways, biomedical research on mammalian systems has revealed over a dozen of oxygenated

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Figure 7 The structure of 13-hydroxyoctadecanoic acid methyl ester and an electron impact mass spectrum of the trimethylsilyl derivative of 13-hydroxyoctadecanoic acid methyl ester synthesized by a microsomal-enriched preparation of Manduca sexta fat body.

metabolites of linoleic acid, some of which exert important biological influence on cells. Products of linoleic acid metabolism include 9-hydroxyoctadecadienoic acid (9(S)-HODE) and 13(S)-HODE. Glasgow and Eling (1990) reported that epidermal growth factor stimulates linoleic acid metabolism in fibroblasts of BALB /c 3T3 mice. The same group later reported that 13(S)-HODE augments the epidermal growth factor receptor signaling pathway (Glasgow et al., 2002). Based on information in the biomedical literature, Putnam and Stanley (unpublished data) considered the hypothesis that insect tissues, also, are able to form potentially bioactive products of linoleic acid. Using fat body from tobacco hornworms, the biosynthesis of two bioactive lipids from radioactive C18:2n
6, specifically 9- and 13-HODE, was recorded. The structures of these compounds were determined by gas chromatography-mass spectrometry (Figure 7). The possibility that metabolites of C18:2n
6 may play important regulatory roles in insect cellular physiology is now emerging. 4.9.6.3. PG Receptors

Turning attention to another emerging topic, recall the broad claim of the eicosanoid hypothesis,

i.e., eicosanoids somehow mediate insect cellular immune reactions to challenge. While the general theme of the hypothesis is strongly supported, information on the mechanisms of eicosanoid actions in insect biology remains the most prominent gap in our current knowledge. In contemporary models of eicosanoid action with respect to immunity (Stanley-Samuelson and Pedibhotla, 1996; Stanley et al., 2002), circulating hemocytes recognize an immune challenge, most likely by recognition of specific cell components, such as LPS in the case of infection with Gram-negative bacteria. Upon stimulation by an invader, hemocytes produce and secrete eicosanoids which enter the hemolymph and influence the action and behavior of other hemocytes. There are important implications in this model. First, the secreted eicosanoids are thought to act on other hemocytes, presumably through functional receptors which have been well characterized in mammalian systems. Second, hemocyte eicosanoid receptors are not uniformly expressed in all hemocytes. To the contrary, because eicosanoids mediate various cell defense actions – such as microaggregation, phagocytosis, cell spreading, nodulation, melanization, and encapsulation – it is possible

332 Eicosanoids

that subpopulations of hemocytes express receptors for particular eicosanoids. The most important implication is that an understanding of eicosanoid receptors can lead to a more detailed understanding of hemocytic immunity and other areas of insect physiology because it will become possible to establish a direct linkage between particular eicosanoids and specific cell actions (Stanley, 2000).

4.9.7. Biochemical Mechanisms of Prostaglandin Actions 4.9.7.1. Background on G Protein-Coupled Receptors

The physiological actions of most PGs are mediated by G protein-coupled receptors (GPCRs) (Breyer et al., 2001). The exceptions are PGA and other cyclopentenone PGs, which readily traverse cellular membranes and interact with intracellular receptors (Negishi et al., 1995b). GPCRs are seven-transmembrane proteins responsible for transducing extracellular signals into intracellular compartments, where cellular responses to the signal are initiated. There are literally hundreds of GPCRs, which are specific for a wide range of ligands, including those involved in vision, olfaction, taste, neurotransmission, neuromodulation, and hormone actions. All multicellular animals have GPCRs, and the genes that encode these receptors make up substantial proportions of their entire genomes. GPRCs make up about 6% of the Caenorhabditis elegans genome, possibly 2% of mammalian genomes, and about 1% of the D. melanogaster genome (Brody and Cravchik, 2000; Vanden Broeck, 2001). The many known GPCRs are sorted into families, including the olfactory receptor family, a glutamate receptor family, a biogenic amine receptor family, and a diuretic hormone receptor family. PGs interact with GPCRs of the rhodopsin family. GPCRs transduce extracellular signals through selective coupling with intracellular G proteins, which in turn regulate the activity of cellular effector proteins, such as adenylyl cyclase, phospholipase C, or ion channels (Blenau and Baumann, 2001). Receptors for most PGs are localized in the plasma membrane, although nuclear PG receptors have been reported (Bhattacharya et al., 1999). PG receptors are classified on the basis of amino acid sequences and the effects of synthetic ligands which interact with the receptors (Coleman et al., 1990). PGs D, E, F, I, and thromboxane A have corresponding receptors, denoted as DP, EP, FP, IP, and TP. So far, four subclasses of PGE receptors have been described,

EP1, EP2, EP3, and EP4. These receptors affect changes in intracellular signal moieties by coupling with various G proteins. For example, EP1 receptors stimulate increased intracellular Ca2þ, and EP2 and EP4 receptors stimulate increased intracellular cAMP. EP3 receptors are unique as so far understood because they exist in variants which differ only in their intracellular C-termini, due to splice variations (Hatae et al., 1989). This allows the different splice variants to couple with multiple G proteins, producing different physiological responses. For example, the EP3A can stimulate increased cAMP concentrations, while EP3B can exert the opposite effect. 4.9.7.2. PG Receptors in Insects

While many GPCRs have been identified and cloned from various insect sources (Roeder, 1999; Blenau and Baumann, 2001; Torfs et al., 2001; Vanden Broeck, 2001), PG receptors have not been reported for insects. Indeed, information on the biochemical and molecular features of insect PG receptors remains the most prominent gap in understanding eicosanoid action in insect physiology (Stanley, 2000). Tunaz and Stanley (unpublished data) now have preliminary data on PGE2 receptors in tobacco hornworm hemocytes. They used classical ligand binding studies, drawn from Negishi et al. (1993) and from Qian et al. (1997), who reported on PGE2 receptors in salivary glands from the lone star tick, to register the presence of PGE2 receptors in membrane preparations made from tobacco hornworm hemocytes. Data from this work indicate the presence of saturable, specific PGE2 receptors associated with membrane preparations of hemocytes from tobacco hornworms. Analysis of the binding data indicted a single binding site model with a KD of approximately 35 nM and Bmax of approximately 7.5 fmol mg
1 protein. It should be borne in mind that analysis of insect PG receptors by radioligand binding studies can be quite problematic. For example, insect tissues are small and they seem to express a small number of PG binding proteins relative to mammalian cells while binding studies require a great deal of tissue. Second, because PGs and other eicosanoids are themselves lipids, they present potentially serious problems with nonspecific or low-affinity binding. Finally, it is very difficult to isolate and study separate subpopulations of insect hemocytes or other subpopulations of cells in other tissues. The difficulties inherent in classical radioligand binding studies have been addressed in work on mammalian PG receptors. The contemporary approach is to clone the genes for the receptors, stably express the genes in established cell lines, and use the cell lines to characterize the receptors. This strategy

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has yielded a very detailed picture of PG receptors in the biomedical context of mammalian systems. Information from the Drosophila and the Anopheles genomes indicates that sequences are present in both insect genomes, which are quite similar to mammalian sequences for PG receptors. In their analysis of the Anopheles genome, Hill et al. (2002) identified a total of 276 GPCRs. Of these, the authors predicted the presence of five classes of PGCRs, including rhodopsin-like receptors. In mammals, PG receptors are members of the rhodopsin-like receptor family. Within the mosquito rhodopsin-like receptors, 22 were identified as ‘‘orphan receptors,’’ that is, receptors for which the physiological ligand is unknown. The predicted number of orphan rhodopsin-like receptors in the Drosophila genome is also 22. Haas and Stanley (unpublished data) used the orphan receptors 1 through 21 from A. gambiae (drawn from the supporting online material in Hill et al. (2002)) to conduct BLAST searches for similar receptor sequences in SWISSPROT. Two of these orphans, GPRorpha5 and GPRorpha11, are very similar to sequences representing receptors for various PGs, with over 100 hits with at least some similarity to mammalian PG receptors and over 40 hits with very high similarity (bit scores >44; E values 115 genes identified by the other studies, Ceriani et al. found that 63 genes remained rhythmic 2 days into DD, and Y. Lin et al. found 22 genes that cycled on the third day of DD. The correlation between more cycles of free-run conditions and fewer identified rhythmic genes is notable. This time-dependent decrement in rhythmicity may reflect a damping effect on molecular rhythms when pacemakers become de-synchronized, as observed in peripheral clocks (Plautz et al., 1997). However, that explanation is not valid in this case as Y. Lin et al. found that all of the canonical clock genes – per, tim, vri, and Clk – continued to cycle robustly 3 days into DD.

The other significant difference between the studies was the treatment of statistics and methods of analysis. To identify rhythmic waveforms, the methods ranged from spectral analysis (Claridge-Chang et al., 2001) to idealized cosine curve fitting (McDonald et al., 2001; Ceriani et al., 2002; Ueda et al., 2002) to direct measurement of intercycle reproducibility of waveforms with autocorrelation analysis (Y. Lin et al., 2002). A concise treatment on the strengths and weaknesses of each strategy was recently detailed by Levine et al. (2002c). It is reasonable to assume that all five studies have correctly identified at least some unambiguous, robustly cycling transcripts. Therefore, until a unified, meta-analysis is performed on the pooled data, a measure of confidence may be afforded by relying upon genes commonly identified across multiple studies. To balance between the degree of agreement amongst the various studies, and the inclusion of the canonical rhythmic genes per, tim, takeout, vri, and Clk, Table 1 lists the 28 genes that were identified by at least three groups to display rhythms in DD. No functional patterns among the known or predicted gene functions seem yet to emerge from the list.

Table 1 The list of Drosophila genes that were commonly identified by at least three microarray studies as circadianly cycling Gene

Location

Known/predicted function

Clk Cyp18a1 Cyp4d21 Cyp6a21 Pdh Slob Ugt35b per tim vri BcDNA:GH02901 CG1441 CG4784 CG4919 CG5156 CG5798 CG5945 CG9645 CG9649 CG10513 CG10553 CG11407 CG11796 CG11853 CG11891 CG14275 CG15093 CG17386

66A12-66A12 17D1-17D1 28A6-28B1 51D2-51D2 72F1-72F1 28B1-28B3 86D5-86D5 3B4-3B4 23F3-23F5 25D4-25D4 13A5-13A5 46C5-46C5 72F1-72F1 94C3-94C3 21F2-21F2 93C1-93C1 34A11-34A11 88B3-88B3 88B3-88B3 96C7-96C7 96C8-96C8 92B3-92B3 77C1-77C1 96C4-96C4 96C6-96C7 29B1-29B1 55F1-55F1 51A1-51A1

RNA polymerase II transcription factor Cytochrome P450, CYP18A1; cytochrome P45; EC:1.14.14.1 Cytochrome P450, CYP4D21; cytochrome P45 Cytochrome P450, CYP6A21; cytochrome P45 Enzyme Signal transduction UDP-glucuronosyltransferase; UDP-glucuronosyltransferase; EC:2.4.1.17 Transcription factor RNA polymerase II transcription factor RNA polymerase II transcription factor Long-chain-fatty-acid-CoA-ligase Cuticle protein; structural protein

Ubiquitin thiolesterase; endopeptidase Endopeptidase Endopeptidase

Luciferase-like; enzyme

3-Hydroxyisobutyrate dehydrogenase-like; enzyme RNA binding

Tick-Talk, the Cellular and Molecular Biology of Drosophila Circadian Rhythms

4.11.4.1.3. The clock also controls the basal level of expression for thousands of genes To control against the false-positive identification of rhythms, all groups performed time-series experiments with arrhythmic flies bearing mutations in clock components such as per01 and ClkJrk. A surprising additional observation from these data was that such mutations produced broad effects on noncycling, basal expression of genes across the Drosophila genome. MacDonald et al., Claridge-Change et al., Ueda et al., and Ceriani et al. observed significant differences in gene expression between ClkJrk mutants and control flies. These changes are likely to represent the absence of normal clock-controlled processes, but may also include gain-of-function consequences that reflect the dominant-negative character of the ClkJrk allele (Allada et al., 2003). Claridge-Change et al. and Y. Lin et al. found that normal expression levels of large swaths of the fly genome depended on per and tim functions. Furthermore, Y. Lin et al. observed that light influenced the expression levels of hundreds of genes when comparing expression levels of control flies under LD versus DD conditions. These light effects were largely dependent upon per function, as per01 mutants did not display comparable changes between LD and DD conditions. Although the power of microarrays to generate candidates for future pursuit cannot be disputed, some caveats should be noted. Several limitations have been enumerated in the reports, such as the heavy dependence of probe sets upon the quality of genome annotation, and the relative insensitivity of microarrays to low-abundance and tissue-specific transcripts. Related to computational annotation is the issue of alternative splicing, a phenomenon thought to contribute heavily to the complexity of multicellular organisms (Harrison et al., 2002). Current generations of microarrays do not yet distinguish between splice variant forms of genes, thus limiting the accuracy of the circadian landscape as seen through a microarray filter. This point was exemplified recently, with the demonstration by Cyran et al. (2003) that only one of six splice variants of the new clock component pdp1 displays transcript rhythms. Is the limited manifestation of transcript rhythms as suggested by Y. Lin et al. and as indicated by the small group of genes coincident between the studies, a meaningful result? One possibility is that despite the existence of a limited number of rhythmic transcripts, a greater population of rhythmic proteins may exist. For example, rhythms in the Lark protein, a protein required for eclosion rhythms, are generated from a nonrhythmic transcript (McNeil

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et al., 1998). Another possibility is that beyond the immediate proximity of the core molecular clock, rhythms exist primarily at the physiological level. Perhaps this is demonstrated by the neurotransmitter PDF, which is neither transcriptionally nor translationally rhythmic, but is rhythmic in its release from axon terminals (J.H. Park et al., 2000; and see below). Likewise, several antecedent studies suggest that even high-amplitude rhythms of transcription in per and tim are not absolutely required for the production of rhythmic behavior (e.g., Frisch et al., 1994; Cheng and Hardin, 1998; Yang and Sehgal, 2001). In sum, these observations all point to the primacy of posttranscriptional events in defining circadian molecular oscillations. Thus, the ‘‘limited manifestation of transcript rhythms’’ may be a reasonable initial description of rhythmicity in the Drosophila transcriptome. Perhaps with future high-throughput techniques such as quantitative mass spectrometry, and the accumulation of more laborious, time-series immunostain-tracking of individual proteins, a broader sense of the Drosophila circadian clock will be achieved. 4.11.4.2. Pacemaker Neurons in the Fly Brain: Anatomy and Roles

4.11.4.2.1. Pacemaker cell types PER expression is found in numerous cells in numerous tissues (Liu et al., 1988, 1992; Siwicki et al., 1988; Zerr et al., 1990; Ewer et al., 1992; Kaneko et al., 1997; Rachidi et al., 1997) and the scale of the pattern depends on the method of visualization. PER immunostaining and per RNA in situ tend to feature small clusters of neurons in the lateral and dorsal brain of the adult, photoreceptors, as well as numerous glia cells in the optic lobes (e.g., Zerr et al., 1990, Ewer et al., 1992, Helfrich-Fo¨ rster, 1995; Kaneko et al., 1997). In contrast, per- and tim-promoter fusions also display these same groups, plus many others not otherwise associated with circadian pacemaking (e.g., Kaneko and Hall, 2000). Price et al. (1998) observed that PER immunostaining was greatly increased in level and distribution (beyond the normal) in a strongly hypomorphic allele of dbt. That result is consistent with the hypothesis that PER is expressed normally, but highly unstable, in a large population of neurons in the larval brain. The canonical PER neuronal groups comprise the lateral cells (LNs) which include the small and large vLNs (s-vLNs and l-vLNs), and dorsal cells (dLNs). There are four to five s-vLNs, of which four express the neuropeptide PDH/PDF (Helfrich-Fo¨ rster, 1995; Kaneko et al., 1997; and see below). There are four to five l-vLNs and approximately seven dLNs.

376 Tick-Talk, the Cellular and Molecular Biology of Drosophila Circadian Rhythms

In the dorsal brain, there are three additional groups: the DN1 cell group contains 10 neurons; the DN2 group is composed of a pair of cells; and the DN3 group contains 30 neurons. The axonal projections of all these neurons appear to target the neuropils of the dorsal protocerebrum, where they intermingle and may interact (Kaneko and Hall, 2000; Helfrich-Fo¨ rster, 2003; Veleri et al., 2003). In addition, a pair of DN1s and some DN3s project to the accessory medulla and appear to provide reciprocal innervation to the s-vLNs. Other DN1s project caudally towards the suboesophageal region and perhaps to the ventral nerve cord.

p0425

4.11.4.2.2. Which PER neurons are necessary for rhythmic behavior? Given the several different pacemaker cell groups, which are necessary and which are sufficient to act as pacemakers to drive rhythmic behavior? Leak and Moore (2000) argue that the different rhythmic systems in mammals are controlled by separate sources of SCN efferents (‘‘core’’ versus ‘‘shell’’). Similar questions can now be applied to the insect brain, where smaller numbers of neurons may facilitate the cellular analysis. Several different genetic designs have tried to define the relative participation of the different pacemaker cells groups in Drosophila. Of these, the study of disconnected mutant flies was among the first and most substantive. The disconnected (disco) gene in Drosophila encodes a widely expressed transcription factor that is required to establish normal sensory connections in the central nervous system (CNS) (Steller et al., 1987; Glossop and Sheperd, 1998). Approximately 5–10% of mutant animals display some retinal-brain connections. While Dushay et al. (1989) found that disco flies are arrhythmic for circadian behaviors, Hardin et al. (1992b) showed that PER levels fluctuate normally in homogenates of disco heads. Those findings suggested that disco acts on the output level because while the mutation eliminated behavioral rhythms, it left the ‘‘clock mechanism’’ intact. An alternative emphasis follows from considering abnormalities of PER expression in disco flies (Helfrich-Fo¨ rster, 1998). PER is found in photoreceptors, DN pacemakers, and putative glia in disco, but is largely absent from LN pacemaker neurons. Blanchardon et al. (2001) subsequently reported that many LNs do in fact survive this mutant background, as indicated by expression of a P{GAL4} reporter line. Together these observations suggest that disco animals retain many functional clock centers (e.g., the retina, the DNs), but display poor behavioral rhythmicity due to the lack of functional clock centers (e.g., LNs) that

normally organize daily locomotion. These animals were very useful to learn specific facts concerning output mechanisms, especially by restricting analysis to mutant animals displaying quasi-normal behavior. Helfrich-Fo¨ rster (1998) monitored the behavior of hundreds of mutant animals: the few that displayed some rhythmic behavior were the only ones to also retain at least one s-vLN. Thus a strong correlation was made between the presence of a s-vLN and circadian locomotor behavior. Mosaic analysis of per expression has also been influential in deciphering the hierarchies of pacemaker cells (Ewer et al., 1992). Frisch et al. (1994) showed that, in complimentary fashion to the disco phenotype, PER expression in just the LNs (both ventral and dorsal) was sufficient to display some rhythmic behavior. Evidence that the DNs also participate in rhythmic output comes from two studies. When PER was returned only to LNs and not to DNs of per mutant flies, the restored rhythms were abnormal in period and strength (Frisch et al., 1994): by inference, DNs may contribute the difference. Similarly, PER expression in accessory medulla neurons under the glass promoter in per mutant flies produced rhythmic behavior (Vosshall and Young, 1995). More recently, Veleri et al. (2003) described a per-luc transgene fusion that restored rhythmicity to per mutant flies under LD cycles, although not under constant conditions. In such flies, the only site of molecular oscillations was in the DN3 populations: strong evidence to suggest these DN neurons are sufficient to provide pacemaker activity for behavior, at least in LD. Peng et al. (2003) and Allada et al. (2003) addressed the same question by asking whether limited expression of transgenic cyc or Clk could rescue either mutant phenotype. Peng et al. (2003) reported that in cycle mutant flies now expressing UAS-cycle under control of pdf-GAL4, PDF neurons alone now displayed proper rhythms of tim RNA, suggesting that UAS-cyc had indeed rescued molecular rhythmic in mosaic fashion. However, the behavior was not rescued, suggesting that other activity in other pacemaker neurons is required. It should be remembered that cyc mutant flies display aberrant PDF neuronal morphology (J.H. Park et al., 2000), and that aberrant PDF neuronal morphology is correlated with arrhythmic behavior (Helfrich-Fo¨ rster, 1998). Allada et al. (2003) used UAS-Clk to rescue two different mutant alleles of Clk. They found that a cry-GAL4 line that is expressed in both vLNs and dLNs (Emery et al., 2000b; but see also Zhao et al., 2003) partially rescued the behavioral rhythmicity of Clk mutants. pdf-GAL4 : UAS-Clk did not provide such rescue. These results, while not yet

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definitive, are consistent with the hypothesis that pacemaker actions of the ventral and dorsal LNs are both necessary and sufficient for Drosophila to display behavioral rhythms. Within the vLN group, five lines of evidence (all indirect) suggest that the s-vLNs may be especially important and the l-vLNs largely unimportant as circadian pacemakers. First, the daily fluctuation in PDF immunostaining of vLN terminals occurs only in the case of the small cell subset (J.H. Park et al., 2000; see further discussion below). Second, the effects of Clock and cyc mutations on pdf RNA levels are seen only in small and not l-vLNs (J.H. Park et al., 2000). Third, in a study of the effects of disco mutations on rhythmicity, Helfrich-Fo¨ rster (1998) confirmed that most or all per-expressing neurons (and hence all pdf vLNs) were typically undetected in disco animals (cf. Blanchardon et al., 2001). She reported a small minority of animals (n ¼ 4) that retained rhythmicity: PDF-expressing s-vLN were visible (as few as one neuron) only in this minority. A l-vLN was also found in a single rhythmic individual, but its processes projected in a s-vLN fashion, to the dorsal protocerebrum. Fourth, Yang and Sehgal (2001) and Shafer et al. (2002) reported fluctuations in PER and TIM immunostaining levels under constant darkness in small, but not in large, vLNs. Finally, cry b mutants fail to maintain rhythms of PER or TIM in DD, in any pacemakers but the s-vLNs (Stanewskey et al., 1998). These diverse and consistent observations form a compelling hypothesis to indicate a special role for the s-vLNs. Direct confirmation of that hypothesis would be a useful step in narrowly defining the pacemaker cell hierarchies within the brain. To do so will require further and more precise manipulation of these neuronal populations. 4.11.4.2.3. PER expression in other insects Several groups have used period gene expression as a method of surveying the number and position of potential circadian pacemakers in other insects. In addition, several of these comparative studies have addressed the potential conservation of circadian pacemaker cell types across insect phylogeny by examining possible coexpression of PER with PDH/PDF. Frisch et al. (1994) used anti-PER antibodies (against a conserved region of the Drosophila PER) to stain the brain of a beetle and reported finding many immunoreactive neuronal groups, some of which had nuclear labeling. In addition to the neurons, they also found many labeled glia in the optic lobes, reminiscent of glial per expression in the Drosophila brain (cf. Ewer et al., 1992).

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Comparisons of anti-PER and anti-PDH stains suggested there was overlap in stained populations, but that many PER-positive did not express PDF. Reppert et al. (1994) cloned a per ortholog from the silkmoth A. pernyii, and reported that antibodies to the moth PER protein labeled the cytoplasm of eight neurosecretory neurons in the protocerebrum (Sauman and Reppert, 1996). Surprisingly, the nuclei of these neurons were never so labeled, the cells were distinct from the PDF-immunoreactive neurons, and no other cells displayed immunostaining. Interestingly, this anti-moth PER antibody when used in Drosophila produced a pattern of PER-like immunoreactivity that was highly similar to that previously found with antibodies to the fly PER (Levine et al., 1995). In addition, the moth protein was functional within Drosophila: it could rescue the per behavioral phenotype when expressed as a transgene in per mutant Drosophila (Levine et al., 1995). The theme of differences between the patterns of PER in Drosophila versus other insects has been repeated in more recent studies, specifically in the moth Manduca (Wise et al., 2002) and the honeybee (Bloch et al., 2003). In both studies, the endogenous PER proteins were used to raise specific antibodies. In Manduca, widespread per expression was found in numerous neurons and glia. Many expressing cells displayed both nuclear and cytoplasmic staining, although evidence for rhythmic expression was only found in glia. Analogous to the observations in A. pernyii, four neurosecretory cells in the pars lateralis of each brain hemisphere exhibited both nuclear and cytoplasmic staining with anti-PER antibodies. These cells were identified as Ia(1) neurosecretory cells that express neuropeptide corazonin immunoreactivity. The accessory medulla contained 100 neurons expressing per RNA but no immunoreactivity. No correspondence of per expression to PDH/PDF expression was evident in any part of the brain. Likewise, in the honeybee brain, PER immunosignals were prominently found in small sets of protocerebral neurosecretory cells, but not within PDH-immunoreactive neurons of the medulla. The latter appeared to have dendrites within the accessory medulla and to project to dorsal protocerebrum, but to lack clock (e.g., period) protein expression. Bloch et al. concluded that: ‘‘. . . although clock proteins are conserved across insect groups, there is no universal pattern of co-expression that allows ready identification of pacemaker neurons within the insect brain.’’ Finally, using an antiDrosophila PER antiserum and anti-PDH, Zavodska et al. (2003) examined several insect

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orders and found consistent expression of both immunosignals in most representatives. However, they reported no correspondence between PER and PDH immunoreactivities. Examples of the lack of nuclear PER staining may reflect a noncircadian function in those cells, or perhaps offer instances when PER’s negative feedback functions may not involve its direct participation in the nucleus (cf. So and Rosbash, 1997). Nevertheless, it remains a challenge for the field to compare the different PER-expressing neuronal populations in different insects, and to rationalize these in terms of neuronal circuits underlying rhythmic behavior. 4.11.4.3. PDF: A Circadian Transmitter

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4.11.4.3.1. Expression Interest in the pigment dispersing factor (PDF) by circadian biologists stems directly from the demonstration of its expression within specific circadian pacemaker neurons of Drosophila (Helfrich-Fo¨ rster, 1995). Its expression within insect tissues, especially in areas associated with circadian pacemakers, was first described several years earlier (Homberg et al., 1991; Na¨ ssel et al., 1991, 1993; Helfrich-Fo¨ rster and Homberg, 1993) using antibodies to crustacean pigment dispersing hormone (PDH, discussed below in Section 4.11.4.3.2). PDH immunosignals are normally found in a limited set of brain and ventral nerve cord cell types. Fundamental elements of the cellular pattern are found throughout insect orders, with salient differences also noted (e.g., Helfrich-Fo¨ rster et al., 1998; Zavodska et al., 2003). Based on its hormonal activities in crustacea (see Chapter 3.16 and see below), there is good evidence to hypothesize that PDF in Drosophila is a secreted neuropeptide and hence a true output signal for certain clock neurons (review: Taghert and Veenstra, 2003). However, recent studies in the cricket indicate that PDH/PDF may also have a nuclear location in some cell types (Chuman et al., 2002). These careful in vivo observations were also supported by studies of PDF transfected into mammalian cells, where the functionality of a putative nuclear localization signal on the PDF precursor was tested. This work underscores the importance of testing all assumptions rigorously and the need to address a range of plausible hypotheses. In Drosophila, PDF expression is limited to the CNS. Within the CNS, there are two PDF cell types in the larva and three cell types in the adult. In larvae, the brain contains four to five LNs that display molecular pacemaker properties (Kaneko et al., 1997). LNs are the likely precursors of the s-vLNs of adults (Helfrich-Fo¨ rster, 1997). In addition, the larval ventral nerve cord contains a

prominent set of four to six large neuroendocrine neurons in the terminal abdominal segments that appear to release PDF into the hemolymph (cf. Persson et al., 2001). The abdominal cells do not display clock properties. In the adult, the vLN group is enlarged by the differentiation of the l-vLNs, which project axons tangentially across a distal layer of the medulla. The abdominal neuroendocrine PDF cell group is maintained. Finally, the third group in adults is a transiently occurring population: a pair of cells in the suboesophageal ganglion of the adult that do not express clock properties and disappear by the second day of adult life. Despite the fact that larvae produce no known circadian output, larval LNs display all the molecular hallmarks of functional pacemakers (e.g., Price et al., 1998). Using anti-PDH (crustacean PDH), an additional two to three cells in the larval protocerebrum and three to five cells in the adult protocerebrum are found. These appear to represent crossreactivity with another, so far unidentified, substance as these cells do not stain with pdf in situ methods (J.H. Park et al., 2000) and they retain immunostaining in pdf mutant animals (Renn et al., 1999). 4.11.4.3.2. The PDH/PDF family of peptides PDH peptides were first studied in crustacea where they cause diurnal movements of pigment granules in retinal cells and their dispersion in epithelial chromatophores. In 1971, a factor that caused the dispersion of distal retinal pigment was purified from eyestalk extracts of the prawn Pandalus borealis (Fernlund, 1971). Upon sequencing, the factor was revealed to be an 18 amino acid peptide with an amidated C-terminus and a free N-terminus (Fernlund, 1976). Originally called light adapting distal retinal pigment hormone (DRPH), it was renamed pigment-dispersing hormone (thus, Panbo-a-PDH), because it also translocates the pigments in the chromatophores centrifugally (Kleinholz, 1975). About a decade later, a second PDH was chemically identified from the eyestalks of the fiddler crab Uca pugilator, a so-called b-PDH that differs from a-PDH in six positions (Rao et al., 1985). To date, PDHs from 15 crustacean species are known (see Chapter 3.16). Extracts from heads of insects were able to elicit a dispersion of pigments in the epidermis of eyestalkless fiddler crabs (Rao et al., 1987). This bioassay then served to isolate the active factor from the grasshopper Romalea microptera, a modified bPDH. Pigment-dispersing factors (PDFs) have since been identified from several insect groups and orthologs identified in the genomes of D. melanogaster

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(Park and Hall, 1998), and in the mosquito A. gambiea (Riehle et al., 2002). The distribution of PDF/PDH appears limited to arthropods to date, but there is one report of immunoreactivity in a mollusk (Elekes and Na¨ ssel, 1999). The PDH / PDF family of peptides displays a large amount of structural conservation; including the length (18 amino acid residues), the N-terminal Asn and C-terminal amidated Ala residues, as well as conserved amino acids at several internal positions. Interestingly, pdh genes in crustaceans have undergone gene duplication and up to three forms, for example, two a-PDHs and one b-PDH, are found in Pandalus jordani (Ohira et al., 2002). Drosophila contains only a single pdf gene and there is no evidence to suggest that PDFs in Drosophila affect pigment dispersion. In various animals including Drosophila, the PDF peptide is predicted to be synthesized following posttranslational processing of a 100 amino acid preproPDF precursor. Nothing is known about the actual biosynthesis of PDF. The general organization of PDF precursors (Ohira et al., 2002) features a signal peptide that is followed immediately by a precursor-associated peptide (PAP) of unknown function, a di- or tri-basic cleavage site and the PDH/PDF octadecapeptide with an N-terminal Gly for amidation, and a mono- or di- or tri-basic cleavage site prior to the translation stop signal. The PAP does not display evolutionary conservation in either its length or primary sequence. 4.11.4.3.3. Cellular release of PDH/PDF There is a rhythm in antibody staining for PDF in the terminals of the s-vLNs of the adult Drosophila brain (J.H. Park et al., 2000 – discussed further below). This observation is thought to reflect a daily rhythm of release. In addition, Kaneko et al. (2000a) and Blanchardon et al. (2001) reported that overexpression of an active tetanus toxin in pdf neurons of Drosophila (with which to cleave synaptobrevin and so reduce evoked release; Sweeney et al., 1995) was not effective in disrupting circadian behavior. It was predicted that disruption of transmitter release by those neurons would have a strong behavioral effect due to results seen when either the pdf gene was mutant or the cells ablated (see below). The lack of a behavioral phenotype in these experiments may be explained by a lack of sensitivity by the PDF peptidergic release system to tetanus toxin. It may be that Drosophila peptidergic release relies on molecules different from synaptobrevin. However, there are two points that suggest caution in accepting that interpretation. For another peptidergic system of Drosophila, release of the eclosion hormone by

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two identified neurons is sensitive to this toxin (McNabb et al., 1997) (see Chapter 3.1). That result suggests at least some Drosophila peptidergic neurons are likely to employ synaptobrevins in the exocytosis of peptidergic secretory granules. In addition, the UAS-tetanus toxin system produces only incomplete cleavage of synaptobrevin (Sweeney et al., 1995), leaving open the possibility that the levels of toxin expression were not sufficient to effect a complete block of release in this instance. 4.11.4.3.4. Control of PDF by the clockworks In mammals, several SCN transmitters display diurnal and/or circadian variation in their expression reflecting control by the clockworks (e.g., Jin et al., 1999). To what extent is PDF expression controlled by clockwork genes? Given a circadian role for PDF signaling, it is now important to ask how the circadian clock produces a diurnal PDF signal. Several cellular phenomena represent potential points of clock regulation: these include pdf transcription and translation, the electrical activity of pdfexpressing neurons, and the sensitivity of PDFreceptive neurons. We now know that PDF expression is regulated by components of the circadian clock, but the details of that regulation are still emerging and they reveal a number of unexpected features. Genetic observations show that pdf RNA is positively regulated by the transcription factors Clock and cyc (J.H. Park et al., 2000) and is also regulated by vrille (Blau and Young, 1999). The effect of the dominant negative ClkJrk allele appeared more severe than that of cyc alleles, consistent with the hypothesis that Clk may have additional partners in mediating its control on pdf (J.H. Park et al., 2000). As yet, no factors have been shown to regulate pdf expression by direct transcriptional activation assays. In the specific case of regulation by CLK : CYC, their actions appear to be indirect (J.H. Park et al., 2000). The regulators Clk and vri produce distinct effects on PDF expression: continuous expression of vri produced a decrease in PDF levels in larval pacemaker neurons (vLNs) but no effect on pdf RNA levels (Blau and Young, 1999). A remarkable feature of this clockwork regulation of PDF expression is its exquisite cell type specificity: it is seen only in clock neurons (not in the abdominal PDF neurons) and, more specifically, only in some clock neurons (the s-vLNs). In spite of its evident clock-controlled transcriptional regulation, pdf RNA does not fluctuate on a daily basis, nor does PDF immunoreactivity vary in large-scale fashion (Park and Hall, 1998). This conclusion was subsequently supported by many microarray experiments (op. cit. Section 4.11.4.1). Clk and cyc

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mutants also displayed aberrations in PDF s-vLN axonal processes, consistent with a normal role in those cells’ morphological differentiation (J.H. Park et al., 2000). Such coordinate control of transmitter expression and axonal morphology by a single transcription factor may reflect a broader developmental theme, akin to similar demonstrations for other nonclock Drosophila neurons (Thor and Thomas, 1997; Allan et al., 2003). In sum, the available data suggest substantial clockwork control of PDF expression. It also indicates that neither pdf transcription nor its translation contribute extensively to the principal gating mechanism within the s-vLN neurons. There is substantial evidence for a daily rhythm of PDF release. In houseflies, PDH injections cause a swelling of L1 and L2 (lamina) axonal diameters that mimics a diurnal change of axon caliber these neurons normally display (Pyza and Meinertzhagen, 1996): an effect consistent with a daytime release of PDF. Drosophila exhibits similar diurnal and circadian changes in laminar axonal diameters and shape (Pyza and Meinertzhagen, 1999). Pyza and Meinertzhagen (1997) also reported that PDH varicosities in the optic lobes display a circadian rhythm in size and spacing: fewer in the subjective day, again consistent with a daily, daytime PDF release. PDF immunoreactivity within s-vLN terminals of Drosophila varies in a diurnal and circadian fashion (J.H. Park et al., 2000). It is high in the subjective day (with a peak 1 h after the start of the subjective day) and low in the subjective night. These findings are also consistent with a hypothesis of release of PDF during the subjective day. This predicted daily PDF release event displays clock influence, in that the period of the staining variation is sensitive to a period length-altering allele of per (J.H. Park et al., 2000). By way of speculation, the daily changes in PDF neurite morphology of the larger flies may find a parallel in the phenotype of the fragile X-related protein gene (dFMR-1: Dockendorf et al., 2002; Inoue et al., 2002; Morales et al., 2002). As discussed below, dFMR-1 mutant animals display both circadian phenotypes and alterations in PDF cell branching. The per and tim mutations did not affect pdf RNA levels, but did affect the level of PDF expression in the s-vLN terminals. Surprisingly, they have opposite effects in this regard. A per0 allele caused consistently low PDF staining, while a tim0 allele caused consistently high PDF staining (J.H. Park et al., 2000). In sum, observations to date indicate multiple levels of control by the clockworks on PDF, specifically within the s-vLNs. Clk and cyc appear to affect pdf gene expression and PDF cell

differentiation, while vri, per, and tim appear to affect a later step(s) in PDF expression, perhaps involving rhythmic transport, processing and/or release. A more complete definition of pdf regulation and an understanding of how its fluctuations contribute to gated PDF signaling represent important future goals. 4.11.4.3.5. PDF physiology and genetics Drosophila mutants for pdf were discovered resident among laboratory stocks of long standing (Renn et al., 1999). These animals contain a nonsense mutation in the signal sequence of preproPDF and are protein nulls. The mutant animals appear and behave normally in most respects. PDF neurons are present and appear fully differentiated in the mutant background. However, the circadian clock-regulated behavior of pdf mutant animals is highly abnormal: while they entrain to light signals, a large majority of the population displays arrhythmicity under constant darkness (DD). The arrhythmicity of pdf mutants is not evident for 1 to 3 cycles of DD. This is unlike the phenotype of animals bearing mutations in clock genes like period or timeless for which arrhythmic behavior is evident as soon as the animal is placed in DD. Transgenic expression of wild-type pdf sequences restored expression of PDF peptide and rhythmic behavior to a great extent. Ablation of the PDF neurons (affected by genetic targeting) in an otherwise wild-type background produced a behavioral phenotype that was in all ways comparable to that produced by pdf mutant flies. A similar conclusion was reached by Blanchardon et al. (2001) who also ablated pdf neurons genetically using a specific GAL4 insertion that prominently features that cell group. Additionally, Helfrich-Fo¨ rster et al. (2000) reported that overexpression of PDF by the UAS : GAL4 system in neurons projecting to the dorsal brain resulted in severe arrhythmicity. That result is consistent with the hypothesis that activation of PDF receptor(s) affects circadian locomotor activity, and that the timing or level of such activation is important to the signaling. Together these observations indicate that the phenotypic deficits exhibited by pdf mutant animals could be attributed in large part to the absence of the pdf gene product, and they lead to several conclusions. First, they contribute directly to the hypothesis that the PDF-expressing vLNs are the primary pacemakers underlying control of daily locomotion (cf. Frisch et al., 1994; Vosshall and Young, 1995). Second, they indicate that pdf is the sole functional output of vLN pacemakers and that PDF is the principal circadian transmitter in

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Drosophila. Third, they predict that other neurons provide auxiliary pacemaking mechanisms and that they must release other (e.g., non-PDF) transmitters (cf. Taghert et al., 2001; Allada et al., 2003; Helfrich-Fo¨ rster, 2003; Peng et al., 2003; Veleri et al., 2003). 4.11.4.3.6. PDF effects on pacemaker synchronization The phenotype of pdf mutant flies is consistent with either of two extreme cellular models. First, pdf signaling may be required to maintain synchronization and/or oscillation among/in pacemaker neurons. Additionally, pdf may be a factor that couples the pacemaker network to premotor centers that govern rhythmic behaviors. These models are not mutually exclusive. Three sets of studies have addressed the first possibility. Petri and Stengl (1997) injected PDH into cockroach brains and reported a significant effect on the phase of subsequent cycles of locomotor activity. PDH caused a phase-dependent 3-h phase delay consistent with it being an input signal that helps sharpen the synchronization of circadian pacemaker neurons. The maximal effect was during the late subjective day, and the shape of the phase-response curve suggested that PDH presents a nonphotic input to the clock, perhaps especially important with respect to coupling the clock outputs from both sides of the brain. In similar fashion, the Drosophila mutant for pdf entrained to LD cycles, but with a phaseadvanced activity peak (Renn et al., 1999). One interpretation of the latter observation is that pdf normally acts to delay the phase of the wild-type activity rhythm. That would be congruent with its pharmacological activity in the larger insects. In this regard, an important role for PDF may therefore be as clock input – to help synchronize the phases of disparate clock neurons. These studies suggest that other transmitters are likely to play similar input roles, promoting either clock phase advances or delays (cf. Volkner et al., 2000; Petri et al., 2002; Wegener et al., 2004). More recently, Peng et al. (2003) analyzed the pattern of tim RNA in situ in the Drosophila brain as a measure of molecular pacemaker function in control versus pdf mutant animals. On DD5, control brains displayed a strong rhythm of tim in situ signals in each of the several sites. In contrast, the amplitude of cycling was much diminished in pdf mutant brains. The authors concluded that pdf was required to maintain high amplitude, molecular rhythms as well as synchronized activity among different pacemaker groups, akin to a recent hypothesis describing the role of neuropeptide VIP receptors in the mammalian SCN (Harmar et al.,

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2002). They speculate that the effect they observe may be related to the demonstration that membrane electrical activity is required by Drosophila pacemaker neurons in order to maintain molecular rhythms (Nitabach et al., 2002). A different conclusion was reached by Blanchardon et al. (2001) who monitored PER immunostaining on DD2-3 in brains of flies that had had the PDF-expressing vLNs (and several other) neurons genetically ablated. That group reported normal PER staining fluctuations in dLN and DN neuronal groups. Two technical differences between these studies are notable and may explain the differences – the amount of time flies were in DD before analysis and the nature of the probes (tim in situ probe versus anti-PER antibodies). 4.11.4.3.7. pdf and geotaxis Toma et al. (2002) reported on studies of highly inbred Drosophila stocks that display either a positive or negative geotaxic bias. Flies called ‘‘Hi’’ typically distribute at higher perches of a vertical maze; the opposite is true of flies called ‘‘Lo.’’ Control animals typically occupy intermediate positions. The authors reasoned that such behavioral biases likely reflect polygenic differences and so they employed microarray technology to determine which genes showed average level differences between the two stocks. Surprisingly, pdf was among the genes found to be reliably different. Further, when tested in the same geotaxic maze device, pdf nulls displayed a negative geotaxic bias as great as that of ‘‘Hi’’ flies. Other clock mutants did not display a similar phenotype. Remarkably, the geotaxic test score was a function of pdf gene dosage; this presumably suggests that the behavior is sensitive to several quantitative levels of PDF peptide. Given the limited sites of PDF expression in the adult animal (essentially two sets of neurons, see above), it should be possible to narrow down the cellular sites of action using molecular genetic techniques. 4.11.4.3.8. PDF receptors There is currently no direct information on the identity of receptors that mediate PDF effects in any organism. Such data would help address many outstanding questions regarding the sites and mechanisms of PDF actions. Peng et al. (2003) reported recently on the distribution of biotinylated PDF in the Drosophila CNS. They applied the labeled probe to fixed brains in whole mount and described a pattern of binding that almost entirely overlapped the pattern of timGAL4 gene expression. That result is consistent with an intriguing hypothesis proposed by those authors – that within the CNS, PDF receptors are

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largely associated with neuronal pacemakers and support their synchronization. However, the report also indicated a need for more characterization of these putative PDF receptor sites: the molecular specificity was uncertain, as the staining effects could not be competed out by unlabeled PDF, even when applied at a 5000-fold excess. In addition, the staining patterns displayed an inconsistent character. Structure-activity studies of the crustacean PDH have revealed possible facts about its interactions with a putative receptor and/or with degradative enzymes (Riehm and Rao, 1982). N-terminal deletions of PDH, removing as many as five amino acids, still resulted in the retention of some (weak) biological activity. Progressively re-extending the molecule back towards the N-terminus restores activity incrementally. Loss of the C-terminal amide resulted in 300-fold loss of activity (Riehm et al., 1985). Because all small neuropeptides act via G proteincoupled receptors (GPCRs; see Chapter 5.5), it is reasonable to think that PDF receptors also act in this manner. Nery and Castrucci (1997) suggested that PDH effects in crustacean chromatophores reflect G’s activation following binding to a GPCR. Hewes and Taghert (2001) found evidence for 44 (1) peptide GPCRs in the Drosophila genome. Those assignments rested on the assumption that Drosophila peptide GPCRs are related to those of other animals by sequence similarities within the highly conserved seven transmembrane domains. Pharmacological study of many of these molecules following functional expression has proceeded using different methods, including signaling assays that rely on a promiscuous G protein (e.g., Cazzamali and Grimmelikhuijzen, 2002; Meeusen et al., 2002), assays of membrane currents in Xenopus (e.g., Park et al., 2002), assays of competitive radioligand binding (e.g., Johnson et al., 2003a), and assays of ligand-dependent translocation of b-arrestin-GFP (Johnson et al., 2003b). To date, roughly 26 of the 44 indicated peptide receptors have been ‘‘de-orphaned’’ by one or more methods. Hopefully, identification and analysis of the PDF receptor(s) will occur in the near future.

cAMP, which in turn activates a cAMP-dependent protein kinase (Nery and Castrucci, 1997) (see also Chapter 3.2). There is genetic evidence supporting the role of the catalytic subunit of protein kinase A (PKA – called DCO) in circadian output controlling locomotor rhythms (Majercak et al., 1997), as well as for a type II PKA regulatory subunit (S.K. Park et al., 2000). However, there is no information yet available to indicate at what cellular/ synaptic level PKA may be acting, nor whether this action reflects PDF action. Finally, Belvin et al. (1999) reported that dCREB2 cycles in circadian fashion, and that it may help sustain rhythmicity of period gene expression by affecting both circadian period and amplitude. These observations suggest that cAMP signaling is likely important at several levels in the circadian system.

4.11.4.4. Second Messenger Systems Mediating PDF and/or Circadian Output

4.11.4.5. Contributions to Circadian Output by Other Transmitters

4.11.4.4.1. cAMP Only a few studies have been conducted on the signaling of PDH in crustaceans. It is suggested that pigment dispersion is achieved by ligand binding to a Gs protein-coupled receptor, resulting in the activation of adenylate cyclase and the increase of the intracellular concentration of

4.11.4.5.1. Other neuropeptides It follows from the chemical heterogeneity of pacemaker neurons (PDF is only expressed by a subset of them) that other transmitters must play important roles in the circuits underlying circadian output. Taghert et al. (2001) presented evidence to suggest that these

4.11.4.4.2. Nf1/MAP kinase Williams et al. (2001) showed an involvement by several putative signaling components that act downstream of PDF in mediating Drosophila circadian output. The Drosophila neurofibromatosis 1 (Nf1) gene product neurofibromin regulates both Ras and cyclic AMP (cAMP) (The et al., 1997). Nf1 mutants exhibited virtually no rhythmic behavior under constant conditions. However, PER and TIM protein, and tim RNA levels in the head continued to oscillate normally in those flies, suggesting a defect downstream of the clockworks. Likewise, PER and TIM proteins also continued to oscillate in the larval LN pacemaker neurons. Furthermore, attempts at phenotypic rescue by restoring wild-type Nf1 only to the pdf-expressing vLN neurons was not effective. Reduced Ras activity partially rescued the circadian rhythm defect of Nf1 mutants, suggesting that NF1 normally regulates Ras in control of this behavior. A link to PDF action was made by demonstrating that phospho-MAP kinase levels were substantially elevated in Nf1 mutants, that such kinase levels cycled diurnally, and that such cycling was reduced in pdf mutants. The authors concluded that PDF driven circadian output is mediated at least in part by NF1 activity through a Ras/MAP kinasedependent signaling pathway (Williams et al., 2001) (see also Chapter 3.2).

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include other amidated peptides different from PDF. In particular, certain peptidylglycine alphahydroxylating monooxygenase (PHM) mosaic animals displayed a more severe locomotor defect than did pdf mutant animals. PHM is an enzyme responsible for the initial step in C-terminal a-amidation, a posttranslational modification displayed by more than 90% of all known Drosophila neuropeptides (Hewes and Taghert, 2001). Importantly, amidation is typically required for full biological activity of neuropeptides (see Riehm et al., 1985, for an example focusing on PDH), and it is specific to secretory peptides. The amidated peptide corazonin is a good candidate for further consideration as it is found in many insects to be co-localized with PER proteins (e.g., Wise et al., 2002; Bloch et al., 2003). Ecdysial behaviors, which are gated by the circadian clock in some cases (e.g., adult eclosion; Truman, 1992) (see also Chapter 3.1), are regulated by several neuropeptides that act in a complex fashion (Ewer and Reynolds, 2002). Release of these factors (including eclosion hormone and CCAP) may be closely (perhaps directly) controlled by clock neurons (Park et al., 2003). 4.11.4.5.2. Other transmitters Two sets of studies highlight circadian changes in the conventional transmitter signaling in the fly. Wide field serotinergic neurons (and PDH-containing neurons) have both been proposed to modulate visual input in the optic ganglia (Na¨ ssel et al., 1991; Pyza and Meinertzhagen, 1997). To investigate possible rhythms in functional aspects of the visual system, Chen et al. (1999) described a diurnal and circadian change in the amplitude of ON and OFF transients in the electroretinogram (ERG) of the fly retina. Consistent with its circadian-regulated release, serotonin injections enhanced the amplitude of ERG transients, and antagonists decreased them. Andretic et al. (1999) found a clock-controlled change in dopamine receptor expression linked to locomotor output (also discussed below in Section 4.11.4.6.5). 4.11.4.6. Evidence for Additional Processes Related to Circadian Output

4.11.4.6.1. The clock controlling rhythmic eclosion Truman and Riddiford (1970) first described the control of adult eclosion by a circadian clock located within the silkmoth brain. They used brain extirpation and reimplantation techniques to demonstrate that an endocrine signal (since identified as the peptide ‘‘eclosion hormone,’’ EH; see Chapter 3.1) is released by the brain according to a signal

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gated by a photoperiod-sensitive clock. Recent studies of the moth have identified specific EH neurons, permitting their comparison to putative sites of period gene expression (Sauman and Reppert, 1996). EH neurons do not appear to be intrinsic circadian pacemakers; instead they are likely driven by higher order pacemakers, which may themselves include other neurosecretory neurons. Myers et al. (2003) re-examined the cellular basis for rhythmic eclosion. They found that circadian oscillations in both the vLNs and the prothoracic gland (PG, a peripheral endocrine organ; see Chapter 3.3) were required for normal rhythmic eclosion. Further, the action of LNs on eclosion and on the rhythms of PER and TIM in the PG was shown to be mediated by its (presumed) secretion of the neuropeptide PDF. The manipulations used affected both clock PDF neurons (vLNs) and nonclock neurons (abdominal neuroendocrine neurons). It is therefore possible that the vLN PDF brain neurons regulate the PG directly (via hemolymph-borne PDF) or indirectly via the neurons themselves controlled by the vLNs. 4.11.4.6.2. A humoral signal controlling daily locomotion Handler and Konopka (1979) performed brain transplants similar to those performed by Truman and Riddiford (1970), but in the much smaller Drosophila to examine the possible influence of brain secretions on rhythmic activity. They found that in a small number of instances, brains from perS animals could restore rhythmicity when transplanted into animals that were otherwise arrhythmic due to their carrying a per 01 allele. The implicated molecule(s) remain unidentified. In summary, there is evidence of diffusible (hormonal) signals normally influencing rhythmic locomotor and eclosion behaviors in insects.

4.11.4.6.3. Fragile X-related protein Several groups have recently implicated the dFMR-1 gene product as a component of circadian output. dFMR1 encodes an RNA-binding protein related to the mammalian fragile X protein. In Drosophila, it is a determinant of neuronal morphology (Zhang et al., 2001; Dockendorf et al., 2002; Morales et al., 2002; Lee et al., 2003; Schenck et al., 2003). Each group determined that dFMR-1 mutant flies were largely arrhythmic in constant darkness (Dockendorf et al., 2002; Inoue et al., 2002; Morales et al., 2002). Inoue et al. (2002) found that the mutants had normal eclosion rhythms, while the other two groups reported either phase (Dockendorf et al., 2002), or phase and amplitude (Morales et al., 2002)

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disturbances in the eclosion rhythm. The vLNs display specific axonal branching defects (Morales et al., 2002), although Dockendorf et al. (2002) reported that these were not observed uniformly. Rhythms of CREB activity were much reduced in mutants (Morales et al., 2002), but rhythmic per expression was maintained (Morales et al., 2002). Interestingly, the double mutant futsch, dFMR-1 suppressed dFMR-1 phenotypes in motor neurons (synaptic overgrowth and defects in neurotransmission; Zhang et al., 2001). However, that genotype did not suppress the dFMR-1 circadian phenotype (Dockendorf et al., 2002). In sum, dFMR-1 mutants maintained clock gene expression, but displayed alterations in behavioral rhythms and in the morphology of pacemaker neurons, and alterations in at least one molecular index of clock output. Determining where and how dFMR-1 acts will provide valuable information concerning which circadian output mechanisms may be defined further. 4.11.4.6.4. takeout Further downstream, the takeout gene contributes to the animals’ starvation responses: this gene responds to starvation with an upregulation of transcription that is dependent on clock gene expression (Sarov-Blat et al., 2000). Its transcription is dependent on expression of most of the major clock genes, but with a distinct delayed phase (So et al., 2000). Additional information concerning takeout functions comes from Dauwalder et al. (2002) who report that the gene is controlled by the sex determination pathway (see Chapter 1.7), and that a loss-of-function allele affects male courtship. Further, they argue that takeout is a member of a large family of genes encoding secreted factors that bind small lipophilic molecules. There exist apparent differences in the reported patterns of takeout gene expression in these various reports. Further studies are needed to help form a consensus regarding takeout functions, and so better understand its role in the circadian system. 4.11.4.6.5. Lark Several lines of evidence demonstrate that the Lark protein participates in construction of a circadian gate for rhythmic eclosion behavior (Newby and Jackson, 1996; McNeil et al., 1998). Lark heterozygotes alter the phase of the eclosion gate, but not its period (Lark homozygotes die during embryogenesis, likely due to loss of other functions). Furthermore, Lark rhythm defects are behavior specific in that they affect eclosion, but not daily locomotor activity. This protein has the molecular signature of an RNA binding protein and is widely expressed in the nucleus of most cells. Significantly, the protein accumulates in the

cytoplasm of specific peptidergic (CCAP) neurons (Zhang et al., 2000). In other insects, these neurons have been implicated in triggering eclosion behavior motor patterns (Ewer and Truman, 1996; Park et al., 2003). Identifying the signal transduction pathways regulated by circadian transmitters and relating Lark actions to the elaboration of a circadian gate are challenges for the near future. 4.11.4.6.6. Circadian regulation of drug sensitivity in Drosophila Hirsh and colleagues have related circadian outputs to drug sensitivity using a decapitated fly assay that allows drug application directly to the CNS. Decapitated flies retain some CNS structures, i.e., the ventral nerve cord, which directly controls somatic musculature and gut motility. Quinpirole, a D2-like dopamine receptor agonist, induces reflexive locomotion, and is most effective during the subjective night (Andretic and Hirsh, 2000). Those studies indicated that dopamine receptor responsiveness is under circadian control and depends on the normal function of the period gene. Flies also show behavioral responsiveness to free base cocaine and sensitization to repeated cocaine doses. Four of five circadian genes tested (per, Clk, cyc, and dbt) altered cocaine sensitization responsiveness (Andretic et al., 1999). Similar phenomena were later described in the mouse (Abarca et al., 2002). Mutant flies also show a lack of tyrosine decarboxylase induction normally seen with cocaine administration. Interestingly, tim01 mutants displayed normal cocaine responses. Andretic et al. (1999) deduce that an unidentified PER binding partner is specifically involved in regulation of drug responsiveness. This result also suggests that drug responsiveness is likely regulated by per expression in a set of cells distinct from those involved in circadian function. 4.11.4.6.7. Circadian output and sleep The biological clock controls many rhythmic output processes, some overt and some obscure. Among these, sleep is arguably the most obvious. For many, it is also the most mysterious. Studies that focus on Drosophila rest as a model began in earnest several years ago (Hendricks et al., 2000; Shaw et al., 2000). Those empirical findings have provided a conceptual framework for which rest in flies can formally correspond to mammalian sleep (reviews: Hendricks, 2003; Shaw, 2003). A significant element in that concept is the demonstration that flies display a robust sleep homeostat – a mechanism to increase the duration of sleep (sleep rebound) within a daily cycle in response to some sleep deprivation experienced in past cycles.

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Given the rhythmic nature of sleep, the possible effects of circadian clock mutations on Drosophila sleep has been an early and enduring point of interest. Two sets of observations describe the current understanding of the relationship between the circadian and sleep mechanisms in Drosophila. First, sleep rebound following a 3-, 6-, or 9-h deprivation protocol is displayed in the cycle immediately following, and it occurs principally during the subjective day (Hendricks et al., 2000; Shaw et al., 2000). Hence, sleep homeostasis is normally gated by the circadian clock. Beyond this permissive feature, does the clock provide regulatory control over sleep mechanisms? The second set of observations suggests the answer is yes. Clock gene mutants display predictable alterations in the dynamics and extent of sleep homeostasis (Shaw et al., 2002; Hendricks et al., 2003). There is clear consensus from observing mutant phenotypes that at least some of the core clock elements may directly regulate aspects of sleep. For example, per and Clk mutants display 100% recovery from sleep deprivation with one daily cycle, whereas control stocks only recover 40% in that same time period. Additionally, cyc and tim mutant flies display increased or decreased amounts of sleep rebound after deprivation that reflect the amount of deprivation experienced. The phenotype of cyc mutant flies in response to sleep deprivation is especially intriguing, and can include exaggerated sleep rebound, increases in baseline sleep amount (set-point), and lethality. That syndrome suggests that not all components of the molecular pathway, as understood in a circadian context, participate equally in a sleep context. Nevertheless, these data together strongly imply that there is a close mechanistic association between elements of the circadian clock and the sleep homeostat. With this strong foundation, we can anticipate that further genetic analysis in Drosophila will make contributions to help define hypotheses by which to describe underlying mechanisms.

Acknowledgments We are very grateful to Erik Johnson, Russ Van Gelder, and Paul Shaw for helpful discussions, and for reading sections of earlier drafts. We apologize to colleagues whose work was not covered for lack of space or oversight. We thank David Van Essen whose pun we refashioned to compose a title for this chapter. YL is supported by the Washington University Medical School MSTP Program. Work in PT’s laboratory is supported by grants from the NINCDS (NS21749) and by the NIMH (MH067122).

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4.12

Insect Transposable Elements

Z Tu, Virginia Tech, Blacksburg, VA, USA ß 2005, Elsevier BV. All Rights Reserved.

4.12.1. Introduction 4.12.2. Classification and Transposition Mechanisms of Eukaryotic TEs 4.12.2.1. Class I RNA-Mediated TEs 4.12.2.2. Class II DNA-Mediated TEs 4.12.2.3. Related Topics 4.12.3. Methods to Uncover and Characterize Insect TEs 4.12.3.1. Early Discoveries and General Criteria 4.12.3.2. Experimental Methods to Isolate and Characterize TEs 4.12.3.3. Computational Approaches to Discover and Analyze TEs 4.12.4. Diversity and Characteristics of Insect TEs 4.12.4.1. Overview 4.12.4.2. RNA-Mediated TEs 4.12.4.3. DNA-Mediated TEs 4.12.5. Search for Active TEs in Insects 4.12.5.1. Identification of Potentially Active TEs on the Basis of Bioinformatics Analysis 4.12.5.2. Detection of TE Transcription 4.12.5.3. Detection of Transposition Events by TE Display 4.12.5.4. Detection of Transposition Events by Inverse PCR 4.12.5.5. Transposition Assay, Reconstruction, and Genetic Screen 4.12.6. Evolution of Insect TEs 4.12.6.1. Genomic Considerations of TE Evolution 4.12.6.2. Vertical Transmission and Horizontal Transfer of Insect TEs 4.12.6.3. Other Possible Evolutionary Strategies 4.12.6.4. Understanding the Intragenomic Diversity of Insect TEs 4.12.7. TEs in Insect Populations 4.12.7.1. Fundamental Questions and Practical Relevance 4.12.7.2. Experimental Approaches 4.12.7.3. Recent Advances 4.12.8. Impact of TEs in Insects 4.12.8.1. TEs and Genome Size and Organization 4.12.8.2. Evolutionary Impact 4.12.9. Applications of Insect TEs 4.12.9.1. Endogenous TEs and Genetic Manipulation of Insects 4.12.9.2. SINE Insertion Polymorphism as Genetic Markers 4.12.9.3. SINE Insertions as Phylogenetic Markers 4.12.10. Summary

4.12.1. Introduction More than half a century ago, Barbara McClintock’s observation of unstable mutations in maize led to the discovery of two mobile genetic elements, Activator (Ac) and Dissociator (Ds) (McClintock, 1948, 1950). The discovery of these mobile segments of DNA, later named as transposable elements (TEs), set forth the revolutionary concept of a fluid and dynamic genome. Five decades later as biology is entering the genomic era, the tremendous diversity of TEs and their potential impact are just being appreciated.

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Being mobile, TEs have the ability to replicate and spread in the genome as primarily ‘‘selfish’’ genetic units (Doolittle and Sapienza, 1980; Orgel and Crick, 1980). They tend to occupy significant portions of the eukaryotic genome. For example, at least 46% of the human genome (Lander et al., 2001) and 16% of the euchromatic portion of the newly reported malaria mosquito genome (Holt et al., 2002) are TE-derived sequences. The relative abundance and diversity of TEs have contributed to the differences in the structure and size of eukaryotic genomes (Kidwell, 2002). Recent evidence suggests

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that the ‘‘selfish’’ property may have enabled TEs to provide the genome with potent agents to generate genetic and genomic plasticity (Kidwell and Lisch, 2000). For example, TEs may have reshaped the human genome by ectopic rearrangements, by creating new genes, and by modifying and shuffling existing genes (Lander et al., 2001). In some cases, TEs had been co-opted to perform critical functions in the biology of their host. One well-documented example is the generation of the extensive array of immunoglobulins and T cell receptors by V(D)J recombination, which is believed to have evolved from an ancient transposition system (Gellert, 2002). Another example is the maintenance of telomeric structures in Drosophila melanogaster by site-specific insertions of two TEs (Pardue and DeBaryshe, 2002). An emerging view is that genomes are dynamic systems, within which diverse TEs evolve. The ‘‘selfish’’ TEs could evolve a wide spectrum of relationships with their hosts, ranging from ‘‘junk parasites’’ to ‘‘molecular symbionts’’ (Brookfield, 1995; Kidwell and Lisch, 2000). The intricate dynamic between TEs and their host genomes is further complicated by the fact that some TEs are capable of crossing species barriers to spread in a new genome. Such a process is referred to as horizontal (or lateral) transfer, which is distinct from the vertical transmission of genetic material from ancestral species/organisms to their offspring. Horizontal transfer may be an important part of the life cycle of some TEs and it may contribute to their continued success during evolution (Silva et al., 2004). From an applied perspective, TEs have been used as tools to genetically manipulate cells/organisms, taking advantage of their ability to integrate cognate DNA in the genome. A well-known example is the transformation system derived from the D. melanogaster P transposable element, which has been instrumental to our understanding of this model genetic organism by providing transformation and mutagenesis tools (see Chapter 4.13). In addition, some TEs have been used as genetic markers for mapping and population studies, taking advantage of their dimorphic insertion states (presence and absence of an insertion) and their interspersed distribution in the genome. For example, the human Alu elements have been shown to be useful population genetic markers (Batzer et al., 1994; Batzer and Deininger, 2002; Salem et al., 2003). The presence and absence of insertions of short interspersed TEs at different genomic loci have also been used as molecular systematics markers to trace the explosive speciation of the cichlid fishes and other vertebrates (Shedlock and Okada, 2000; Terai et al., 2003).

The focus of this chapter is on recent advances in the study of insect TEs. A brief introduction on TE classification and transposition mechanisms will be provided, followed by sections that describe the current approaches to study insect TEs and sections that highlight the diversity and evolutionary dynamics of TEs in insect genomes. Applications of TEs in genetic and molecular analysis of insects will be discussed in the end. Readers may consult recent reviews (Silva et al., 2004) (see Chapter 4.13) and the second edition of a book on mobile DNA (Craig et al., 2002) for details on related topics.

4.12.2. Classification and Transposition Mechanisms of Eukaryotic TEs TEs can be categorized as Class I RNA-mediated or Class II DNA-mediated elements according to their transposition mechanisms (Finnegan, 1992). The transposition of RNA-mediated TEs involves a reverse transcription step, which generates cDNA from RNA molecules (Eickbush and Malik, 2002). The cDNA molecules are integrated in the genome, allowing replicative amplification. The transposition of DNA-mediated elements is directly from DNA to DNA and does not involve an RNA intermediate (Craig, 2002). In most cases, both classes of TEs will create target site duplication (TSD) upon their insertion in the genome (Figure 1). Both classes can be further categorized into different groups and all groups of TEs discussed here have been found in various species of insects. There have been several recent reviews on different groups of TEs in both classes (Deininger and Roy-Engel, 2002; Eickbush and Malik, 2002; Feschotte et al., 2002; Robertson, 2002). 4.12.2.1. Class I RNA-Mediated TEs

RNA-mediated TEs include long terminal repeat (LTR) retrotransposons, non-LTR retrotransposons, and short interspersed repetitive/nuclear elements (SINEs). Non-LTR retrotransposons are also referred to as retroposons or long interspersed repetitive/nuclear elements (LINEs). The structural features of the three groups of RNA-mediated TEs are illustrated in Figure 2 using representatives from different insects. All RNA-mediated TEs produce RNA transcripts that are reverse transcribed into cDNA to be integrated in the genome (Eickbush and Malik, 2002). Detailed mechanisms used by LTR and non-LTR retrotransposons are elegantly described in two recent reviews (Eickbush, 2002; Voytas and Boeke, 2002).

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Figure 1 Mechanism of generating target site duplication (TSD). The TSDs, which are marked by open arrows, are not part of the TE sequence. They are target sequences duplicated upon a TE insertion. Most TEs create TSDs although the Helitron transposons and some non-LTR retrotransposons do not. See recent reviews (Kapitonov and Jurka, 2001; Craig, 2002; Eickbush and Malik, 2002) for illustrations of the transposition mechanisms of different types of TEs.

4.12.2.1.1. LTR retrotransposons LTR retrotransposons transpose through a mechanism much like that used by retroviruses. The LTRs in the LTR retrotransposons are generally 200 to 500 bp long and are involved in all aspects of their life cycle that includes providing promoter sequences and transcription termination signals (Eickbush and Malik, 2002). As shown in Figure 2, LTR retrotransposons encode a pol(polymerase)-like protein that contains reverse transcriptase (RT), ribonuclease H (RNase H), protease (PR), and integrase (IN) domains that are important for their retrotransposition. The RT domain performs the key function of reverse transcription and its sequence has been used for phylogenetic classification of LTR retrotransposons into four clades including Ty1/copia, Ty3/ gypsy, BEL, and DIRS (Eickbush and Malik, 2002). The IN domain is responsible for inserting the cDNA copy into the host genome. In addition to the pol-like protein, LTR retrotransposons encode an additional protein related to the retroviral gag (group-associated antigene or group-specific antigen) protein that binds nucleic acid or forms the nucleocapsid shell. Some LTR retrotransposons also have an env(envelope)-like fragment that encodes a transmembrane receptor-binding protein that allows the transmission of retroviruses. Some of the LTR retrotransposons that encode an env protein are in fact retroviruses (Eickbush and Malik, 2002). 4.12.2.1.2. Non-LTR retrotransposons Non-LTR retrotransposons, or LINEs, or retroposons are generally 3 to 8 kb long and have been found in virtually all eukaryotes studied. Like the LTR retrotransposons, most non-LTR retrotransposons also have a pol-like protein that includes a RT domain that

is essential for their retrotransposition. The RT domain has been used for phylogenetic classification of non-LTR retrotransposons into 17 clades, most of which probably date back to the Precambrian era, approximately 600 million years ago (Malik et al., 1999; Eickbush and Malik, 2002; Biedler and Tu, 2003). Some elements also have an RNase H and/or AP endonuclease (APE) domain encoded in the pol-like open reading frame (ORF). In addition to the pol-like protein, many non-LTR retrotransposons encode a protein that is related to the retroviral gag protein. Studies of a gag-like protein from L1 retrotransposon in mice show that it acts as a nucleic acid chaperone (Martin and Bushman, 2001). Other typical structural characteristics found in various non-LTR retrotransposons are internal Pol II promoters and 30 ends containing AATAAA polyadenylation signals, poly(A) tails, or simple tandem repeats. Target primed reverse transcription has been proposed as the mechanism of retrotransposition for R2 of Bombyx mori and may be generally true for all non-LTR elements (Luan et al., 1993; Eickbush, 2002). Because they transpose by target primed reverse transcription, some non-LTR retrotransposons could rely rather heavily on host DNA repair mechanisms, and this relationship with the host may give non-LTR retrotransposons some flexibility with regard to the domains required in an autonomous element (Eickbush and Malik, 2002). Some non-LTR retrotransposons such as R2 are site-specific because their endonucleases make precise cleavage at specific targets (Xiong and Eickbush, 1988; Eickbush, 2002). 4.12.2.1.3. SINEs SINEs are generally between 100 and 500 bp long and they do not have any coding potential. SINEs may have been borrowing the

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Figure 2 Structural characteristics of representative Class I RNA-mediated TEs in insects. Representatives are shown from three major groups: (a) long terminal repeat (LTR) retrotransposons, (b) non-LTR retrotransposons, and (c) short interspersed repetitive elements (SINEs). The name of each representative element, its host species, and its approximate length are shown as the heading. Open reading frames (ORFs) are shown as open boxes. The elements are not drawn to scale. References for information on these RNA-mediated elements are the following. BEL1, Davis and Judd (1995); Feilai, Tu (1999); Gypsy, Mizrokhi and Mazo (1991); I, Fawcett et al. (1986); L1, Biedler and Tu (2003); Outcast, Biedler and Tu (2003). env, envelope protein; gag, group-associated antigene or group-specific antigen; pol, polymerase-like protein; APE, apurinic-apyrimidinic endonuclease; IN, integrase; LINEs, long interspersed repetitive elements; LTR, long terminal repeat; PR, protease; RH, RNase H; RT, reverse transcriptase.

retrotransposition machinery from autonomous non-LTR retrotransposons, which may be facilitated by similar sequences or structures at the 30 ends of a SINE and its ‘‘partner’’ non-LTR retrotransposon (Ohshima et al., 1996; Okada and Hamada, 1997). Unlike non-LTR retrotransposons that use internal Pol II promoters, SINE transcription is directed from their own Pol III promoters that are similar to those found in small RNA genes. SINEs can be further divided into three groups based on similarities of their 50 sequences to different types of small RNA genes. Elements such as the primate Alu family share sequence similarities with 7SL RNA (Jurka, 1995)

while most other SINEs belong to a different group that shares sequence similarities to tRNA molecules (Adams et al., 1986; Okada, 1991; Tu, 1999). Recently, a new group of SINEs named SINE3 have been discovered in the zebrafish genome, which share similarities to 5S rRNA (Kapitonov and Jurka, 2003a). Some non-LTR retrotransposons tend to generate truncated copies due to incomplete reverse transcription during cDNA synthesis. Although these short copies of RNA-mediated TEs are also called SINEs (Malik and Eickbush, 1998), they should not be confused with the true SINEs that use Pol III promoters.

Insect Transposable Elements

4.12.2.2. Class II DNA-Mediated TEs

DNA-mediated TEs include cut-and-paste DNA transposons (Figure 3), miniature inverted-repeat TEs (MITEs) (Figure 4), and a recently discovered group called Helitrons (Kapitonov and Jurka, 2001, 2003b). Their transposition is directly from DNA to DNA, and does not involve an RNA intermediate (Craig, 2002). 4.12.2.2.1. Cut-and-paste DNA transposons DNA transposons such as P, hobo, and mariner are usually characterized by 10–200 bp terminal inverted repeats (TIRs) flanking one or more ORFs that encode a transposase. They usually transpose by a cut-and-paste mechanism and their copy number can be increased through a repair mechanism (Finnegan, 1992; Craig, 2002). As shown in Figure 3, cut-and-paste DNA transposons can be subdivided into several families or superfamiles according to their transposase sequences and these families/ superfamilies are also characterized by TSDs of specific sequence or length. The families/superfamilies that have been found in insects include IS630Tc1-mariner, hAT, piggyBac, PIF/Harbinger, P, and Transib (Shao and Tu, 2001; Robertson, 2002; Kapitonov and Jurka, 2003b). 4.12.2.2.2. MITEs MITEs are widely distributed in plants, vertebrates, and invertebrates (Oosumi et al., 1995; Wessler et al., 1995; Smit and Riggs, 1996; Tu, 1997, 2001a). Most MITEs share common structural characteristics such as TIRs, small size, lack of coding potential, AT richness, and the potential to form stable secondary structures (Wessler et al., 1995). MITEs may have been ‘‘borrowing’’ the transposition machinery of autonomous DNA transposons by taking advantage of shared TIRs (MacRae and Clegg, 1992; Feschotte and Mouches, 2000b; Zhang et al., 2001). However, an alternative hypothesis suggests that they may transpose by a hairpin DNA intermediate produced from the folding back of single-stranded DNA during replication, which may better explain how MITEs could achieve immensely high copy numbers in some genomes (Izsvak et al., 1999). The evolutionary origin of MITEs is still not clear (see Section 4.12.4.3.3). One obvious source of MITEs would be internal deleted autonomous DNA transposons (Feschotte and Mouches, 2000b). In this case, MITEs are basically nonautonomous deletion derivatives of DNA transposons. Recent studies show that the similarities between many MITEs and their putative autonomous partners are restricted to the TIRs (Feschotte et al., 2003)

399

(Figure 4). Although subsequent loss of autonomous partners in the genome remains a possible explanation for the lack of internal sequence similarity between MITEs and autonomous DNA transposons, two other explanations are perhaps more plausible. First, MITEs could originate de novo from chance mutation or recombination events resulting in the association of TIRs flanking unrelated segments of DNA (MacRae and Clegg, 1992; Tu, 2000; Feschotte et al., 2003). Alternatively, MITEs could originate from abortive gap-repair following the transposition of DNA transposons, which has been shown to occasionally introduce transposonunrelated sequences (Rubin and Levy, 1997). 4.12.2.2.3. Helitrons: the rolling-circle DNA transposons Helitrons and related transposons have recently been discovered in insects and plants, which appear to use a rolling-circle mechanism of transposition (Le et al., 2000; Kapitonov and Jurka, 2001; 2003b). Instead of cut-and-paste transposase, Helitrons encode proteins similar to helicase, ssDNA-binding protein, and replication initiation protein. These proteins facilitate the rolling-circle replication of Helitrons, a mechanism previously described for the bacterial IS91 transposons (Garcillan-Barcia et al., 2002). 4.12.2.3. Related Topics

4.12.2.3.1. Foldback elements Drosophila Foldback elements are characterized by very long inverted repeats (Truett et al., 1981). It is not known how Foldback elements transpose, although the presence of long inverted repeats indicates a possible DNA-mediated mechanism. Some researchers group Foldback elements as a distinct class, namely Class III (Kaminker et al., 2002). 4.12.2.3.2. What is a family? Before we move ahead with discussions on the discovery and diversity of insect TEs in the next two sections, it may be helpful to clarify the use of the term ‘‘family’’ in the context of TEs. The term ‘‘family’’ is often used to refer to a group of related TEs in diverse organisms that usually share conserved amino acid sequences in their transposase or RT and one such example is the mariner family. A TE also consists of many copies generated by transposition events in a genome, and therefore these related copies are sometimes also referred to as a family. Some families consist of multiple distinct groups that are subdivided into subfamilies. Obviously, relatedness is a relative concept in evolution and thus a working definition is needed in each case until a universal family definition is developed.

p0080

400 Insect Transposable Elements

Figure 3 Structural characteristics of representative cut-and-paste DNA transposons in insects. The name of each representative transposon, its host species, and its approximate length are shown as the heading of each panel. Open arrows indicate target site duplications (TSDs). Filled triangles indicate terminal and subterminal inverted repeats. The lengths of these inverted repeats are marked. Exons are shown as open boxes and introns are shown as filled black boxes. The 50 and 30 untranslated regions are not shown. The elements are not drawn to scale. References for information on these transposons are the following: P, Engels (1989) and Rio (2002); hobo, Streck et al. (1986); mariner Mos1, Jacobson et al. (1986); Minos, Franz and Savakis (1991); ITmD37E, Shao and Tu (2001); Pogo, Tudor et al. (1992) and Feschotte and Mouches (2000b); piggyBac, Cary et al. (1989); PIF/Harbinger-like, Biedler et al. unpublished data; Transib1, Kapitonov and Jurka (2003b).

Insect Transposable Elements

Figure 4 Relationship between PIF/Harbinger-like DNA transposons and related MITEs in Anopheles gambiae. (a) Structural features of PIF/Harbinger-like DNA transposons. There are multiple PIF/Harbinger-like DNA transposon families in A. gambiae, all of which have the characteristic AT-rich 3 bp TSDs (Biedler et al., unpublished data). Although the TIRs are highly conserved between different copies of a family, they are variable between different PIF/Harbinger-like transposon families. (b) Structural features of deletion derivatives of PIF/Harbingerlike DNA transposons. Only two elements are apparent deletion derivatives of two different PIF/Harbinger-like transposons. (c) Structural features of PIF/Harbinger-related MITEs. These putative MITEs share similar TSDs and TIRs with PIF/Harbinger-like DNA transposons, although there are no internal sequence similarities between them. Twenty families of PIF/ Harbinger-realted MITEs belong to this category. Note that the term ‘‘family’’ is used here to refer to a group of similar copies.

4.12.3. Methods to Uncover and Characterize Insect TEs 4.12.3.1. Early Discoveries and General Criteria

Before the availability of the large amount of genomic sequence data, TEs were often discovered by serendipitous observations during genetic experiments. As described above (see Section 4.12.1), McClintock’s observation of the unstable mutations in maize led to the discovery of two mobile genetic elements Ac and Ds, although the molecular characterization of these elements came many years later (review: Fedoroff, 1989). Similarly, the observation of an unstable white-peach eye-color mutation in Drosophila mauritiana led to the discovery of the mariner transposon (Hartl, 1989). The piggyBac transposon was discovered as an insertion in a baculovirus after passage through a cell line of the cabbage looper Trichoplusia ni (review: Fraser, 2000). In a slightly different vein, the D. melanogaster P and I elements were discovered because of their association with a genetic phenomenon called hybrid dysgenesis, which refers to a group of abnormal traits including high mutation rates and sterility in crosses of certain strains (Kidwell, 1977; Finnegan, 1989). The genetic mutations described above, albeit rare, tend to identify active transposition events that resulted from active TEs in the genome.

401

The repetitive nature of TEs can also be used for their discovery and isolation, although not all repetitive elements are TEs. When DNA sequences are available, TEs can be identified on the basis of either similarity to known TEs, or common structural characteristics. In some cases, evidence for past TE insertion events could be identified on the basis of sequence analysis, which further supports the mobile history of a particular element. Although the criteria and methods described in this section are not unique to insects, it may be necessary to visit this topic here because of the lack of a systematic review on these issues and because of the growing interests in TE analysis in the current genomic environment. 4.12.3.2. Experimental Methods to Isolate and Characterize TEs

Several experimental methods have been used to discover TEs on the basis of their repetitive nature. Although relatively straightforward, these methods may not clearly distinguish between TEs and other repetitive sequences in the genome. In other words, the repeats discovered using these methods are not always TEs. One way to discover repeats in the genome is to isolate visible bands in an agarose gel running a sample of restriction enzyme-digested genomic DNA. This is based on the assumption that only highly reiterated sequences containing two or more conserved recognition sites for the restriction enzyme will produce a visible band among the smear of digested genomic DNA, and the bands can be cut out from the gel and purified for cloning and sequencing. Another approach to search for repeats is to screen a genomic library using labeled genomic DNA as a probe. This approach can be used effectively to identify abundant or highly repetitive sequences in the genome, which is based on the principle that only the repetitive fraction of the genome will produce a sufficient amount of labeled fragments that will generate hybridization signals during the screening (Gale, 1987; Cockburn and Mitchell, 1989). A third method is to use Cot analysis to help identify repetitive sequences in the genome, which is based on DNA reassociation kinetics (Adams et al., 1986; Peterson et al., 2002). For example, Cot analysis of genomic DNA can be performed to isolate the moderately repetitive portion of the genome that tends to contain TEs, and one can construct a subgenomic library using this fraction of genomic DNA to search for potential TEs. Several methods can be used to identify and isolate TEs on the basis of information derived from related TEs. For example, homologous TE probes may be used in Southern blotting and genomic library

402 Insect Transposable Elements

screening experiments to identify related TEs. Polymerase chain reaction (PCR) analysis using primers that are conserved between related TEs can also be used to isolate different members of a TE family. 4.12.3.3. Computational Approaches to Discover and Analyze TEs

Insect science is rapidly entering a new era as demonstrated by the publication of the D. melanogaster genome sequence (Adams et al., 2000), the report of the genome assembly of the African malaria mosquito Anopheles gambiae (Holt et al., 2002), and the recent progress in genome projects of a number of other insects including the yellow fever mosquito Aedes aegypti, the honeybee Apis mellifera, the silkworm Bombyx mori, and the tobacco budworm Heliothis virescens (Kaufman et al., 2002; Transgenesis and Genomics of Invertebrate Organisms, 2003). This genome revolution is producing an ever-expanding sea of data that can be explored using a bioinformatics approach to identify interspersed TEs. As described below, a few new tools have been developed which represent a shift from merely masking TEs (e.g., RepeatMasker; review: Jurka, 2000) to the discovery, annotation, and genomic analysis of TEs. The use of bioinformatics tools provides great advantages by allowing analysis of TEs in the entire genome and by allowing quick surveys of a large number of TE families to identify the most promising candidates for discovering active TEs (see Section 4.12.5) and for population analysis (see Section 4.12.9). It should be noted that these approaches are not limited to fully sequenced genomes. Because of the repetitive nature of TEs, sequences from a small fraction of a genome tend to contain a large number of TEs that may be discovered using the bioinformatics approaches described here. Of course, greater numbers of sequences and longer assembly would be beneficial in analyzing low copy number or long TE sequences. It should also be mentioned that no high-end computing facilities are required for the majority of these bioinformatics programs. 4.12.3.3.1. Homology-dependent approaches Searching for TEs in a genome on the basis of similarities to known elements discovered in different species is relatively straightforward. However, given the diversity and abundance of TEs, systematic computational approaches are necessary for efficient and comprehensive analysis. One such program was reported that uses profile hidden Markov models to find all sequences matching the full-length RT with the conserved FYXDD motif common to all reverse transcriptases (Berezikov

et al., 2000). A BLAST-based systematic approach to simultaneously identify and classify TEs is developed recently (Figure 5). This approach incorporates multiquery BLAST (Altschul et al., 1997) and a few computer program modules (freely available at jaketu.biochem.vt.edu) that organize BLAST output, retrieve sequence fragment, and mask database for identified TEs. The method was successfully used to discover and characterize non-LTR retrotransposons in the A. gambiae genome assembly (Biedler and Tu, 2003). The strategy is explained using the example of a representative non-LTR retrotransposon as shown in Figure 5a. The potentially comprehensive nature of this approach was demonstrated as two new clades were identified (Biedler and Tu, 2003) (see Section 4.12.4). The inclusive nature of this approach was further indicated when observed that non-LTRs across all existing clades were identified using the reiterative approach with a single D. melanogaster representative in the Jockey clade as the starting query. The reiterative feature of the strategy described in this study provides an opportunity for further automation by linking the modules described in Figure 5a. An early version of a fully automated program was tested, which was named TEpipe, to identify all non-LTR retrotransposon families in the A. gambiae genome. Preliminary results showed that 102 of the 104 non-LTR retrotransposon families were identified in one run of the program which takes less than 2 h on a Linux workstation. The robustness of the family classification assigned by the program was confirmed using a few independent tests. Alignment of the nucleotide sequences plus flanking genomic sequence of each family was performed with ClustalW (Thompson et al., 1994) to determine transposon boundaries, full-length elements, and TSDs (Figure 5b). The alignment process has recently been automated. The TEpipe approach should work for any TE with coding capacity in any genome. 4.12.3.3.2. Homology-independent approaches A suite of computer programs has been developed recently to search large databases rapidly for sequences with characteristics of MITEs (Tu, 2001a). The key program, FINDMITE1, searches the database for inverted repeats flanked by userdefined direct repeats within a specified distance range (Figure 6). The program uses the idea of the Knuth–Morris–Pratt string-matching algorithm (Tu, 2001a) to speed up the pattern match shifts. FINDMITE1 was used to uncover eight novel families of MITEs in A. gambiae (Tu, 2001a). Improvements have been made so that the new version of

Insect Transposable Elements

403

Figure 5 Design of TEpipe, a pipeline program for simultaneous identification and classification of TEs on the basis of sequence similarities. The example shown here is the strategy used to analyze all non-LTR retrotransposons in the Anopheles gambiae genome (Biedler and Tu, 2003). (a) Strategy to identify non-LTR retrotransposon families. (b) Strategy to define full-length elements. Ovals indicate databases used for searches. Rectangles indicate input/output files. Program modules are in bold beside arrows. Modules written in our laboratory are available at the website jaketu.biochem.vt.edu. (Biedler and Tu, 2003). TSDs, target site duplications. (Reproduced with permission from Biedler, J., Tu, Z., 2003. Non-LTR retrotransposons in the African malaria mosquito, Anopheles gambiae: unprecedented diversity and evidence of recent activity. Mol. Biol. Evol. 20, 1811–1825; ß Oxford University Press.)

FindMITE handles whole-genome sequence databases better, incorporates downstream analysis, and produces fewer false positive results which could be overwhelming in some cases. A program module AlignMITE has been developed to identify candidate MITEs that share the same TIRs and automatically align them by calling ClustalW (Thompson et al., 1994) to determine MITE boundaries and full-length elements. Another program module, MITEInsertion, has been developed to uncover evidence of MITE insertion. Because a

whole-genome database is not required, this systematic approach could have broad applications for the analysis of the model genomes as well as the vast majority of the less sequenced genomes. Although FINDMITE1 was originally designed to discover and analyze MITEs, it is possible to use it for the identification and characterization of DNA transposons that also contain TIRs. It is possible to use TEpipe to survey for the transposase-encoding sequences and use FINDMITE to analyze the TIRs and TSDs.

404 Insect Transposable Elements

Figure 6 Design of two programs to search for different groups of TEs on the basis of shared structural features. (a) FINDMITE1 is a C program designed to rapidly search a database for sequences that have the characteristics of MITEs (Tu, 2001a). The program searches sequences in the database for inverted repeats flanked by user-defined direct repeats within a specified distance range. FINDMITE1 is available at our website (jaketu.biochem.vt.edu). The newly developed MITEpipe program, which further reduces false positive signals and incorporates FINDMITE1 with downstream analysis, will be available soon at the same website. (b) LTR_STRUC is a program that identifies LTR retrotransposons on the basis of the presence of long terminal repeats (most LTRs contain TG. . .CA termini), target site duplications (TSDs), and additional information such as primer binding site and polypurine tract (McCarthy and McDonald, 2003). The program is available as a console application from its authors.

LTR_STRUC is a program that identifies LTR retrotransposons on the basis of the presence of LTRs (most LTRs contain TG. . .CA termini), TSDs, and additional information such as primer binding site and polypurine tract (McCarthy and McDonald, 2003) (Figure 6b). Although it is not designed to uncover solo LTRs or truncated LTR retrotransposons, the program offers a rapid and efficient approach to systematically identify and characterize LTR retrotransposons in a given genome and it can be used as a discovery tool for new families of LTR retrotransposons. The program is available as a console application from its authors. Recently, an automated program named RECON has been reported (Bao and Eddy, 2002), which identifies TE sequences on the basis of their repetitive nature in the genome. RECON uses a multiple sequence alignment algorithm which represents a significant improvement to previous methods based on the same strategy. Because it does not rely on sequence homology or structural information, RECON is potentially comprehensive and may be able to identify all repetitive sequences. By the same token, RECON is computationally intensive as the size of the genome database increases. A two-step approach in which a large number of repeats are identified and masked first using a small fraction of the genome sequence may resolve this potential problem.

copia, gypsy, I, R1, P, mariner, hobo, piggyBac, and transib are the founding members of several diverse families/superfamilies that were later shown to have broad distributions in eukaryotes. In addition, recent studies revealed a few novel and intriguing TEs in insects that are described in detail below. Tables 1 and 2 provide a relatively extensive, but by no means exhaustive, compilation of the two classes of TEs in insects. Figures 2 and 3 depict the characteristics of representatives of different RNA-mediated TEs and the cut-and-paste DNA transposons. In addition to the two ‘‘fully sequenced’’ dipterans, a few other dipteran species and a lepidopteran species (Bombyx mori) represent the sources from which the majority of TEs have been discovered and analyzed. As more insect genome projects move ahead and studies on insect TEs expand, more TEs from diverse species will be discovered which will expand our horizon and offer new insights from a comparative genomics perspective. In this section, a detailed account of the characteristics of all different families of TEs will not be discussed. Instead the focus will be on recent advances and interesting features of some novel insect TEs. In anticipation of the explosion of TE discovery in the near future, my laboratory will periodically upload updated Tables 1 and 2 on a website (jaketu. biochem.vt.edu). 4.12.4.2. RNA-Mediated TEs

4.12.4. Diversity and Characteristics of Insect TEs 4.12.4.1. Overview

Virtually all classes and types of eukaryotic TEs have been found in insects. Insect TEs such as

4.12.4.2.1. LTR retrotransposons The distribution of LTR retrotransposons in insects is shown in Table 1. The structural characteristics of representatives of three major groups found in insects, Ty1/copia, Ty3/gypsy, and BEL, are shown in Figure 2. In addition to the LTRs that contain the

Insect Transposable Elements

t0005

405

Table 1 RNA-mediated transposable elements in insects Superfamily (clade)

Element and referenceb

Order

Organismc,d

Diptera

Aedes aegypti

Diptera Diptera Lepidoptera Coleoptera Diptera Diptera Diptera Diptera

Anopheles gambiae Drosophila melanogaster Bombyx mori Tribolium castaneum Anopheles gambiae Anopheles gambiae Ceratitis capitata Drosophila melanogaster

Lepidoptera Lepidoptera Lepidoptera Lepidoptera Diptera Diptera Diptera Lepidoptera Lepidoptera

Bombyx mandarina Bombyx mori Lymantria dispar Trichoplusia ni Aedes aegypti Anopheles gambiae Drosophila melanogaster Bombyx mandarina Bombyx mori

I. LTR retrotransposonsa Ty1/copia

Ty3/gypsy

BEL

Mosqcopia (Tu, unpublished data), Zebedee (Warren et al., 1997) copia-like (Holt et al., 2002), mtanga (Rohr et al., 2002) 1731, copia, Dm88, frogger Yokozuna (Ohbayashi et al., 1998) Woot (Beeman et al., 1996) Beagle, gypsy-like, Cruiser, Osvaldo, Springer (Holt et al., 2002) Ozymandias (Hill et al., 2001) yoyo (Zhou and Haymer, 1998) 17.6, 297, 412, accord, blastopia, blodd, Burdock, Circe, gtwin, gypsy, HMS Beagle, Idefix, invader, McClintock, mdg1, mag3, micropia, opus, qbert, Quasimodo, rover, springer, Stalker, Tabor, Tirant, Transpac, Zam Judo, Karate (Abe et al., 2002) Kabuki (Abe et al., 2000), Mag (Garel et al., 1994) Lydia (Pfeifer et al., 2000) TED (Friesen and Nissen, 1990) MosqNinja (Tu, unpublished data) Moose, Pao-like (Holt et al., 2002) aurora, Bel, diver, GATE, roo, rooA Yamato (Abe et al., 2002) Pao (Xiong et al., 1993), Kamikaze, Yamato (Abe et al., 2001)

II. Non-LTR retrotransposonsa R4 R2 L1 L2 RTE R1

LOA

I

Jockey

CR1 Loner Outcast

Not classified

III. SINEs tRNA-related SINEs

Ag-R4_ 1 (Biedler and Tu, 2003) Dong (Xiong and Eickbush, 1993) R2 (Eickbush and Malik, 2002) R2Bm (Xiong and Eickbush, 1988) Ag-L1_ 1–5 (Biedler and Tu, 2003) Ag-L2_ 1–3 (Biedler and Tu, 2003) JAM1 (GenBank Z86117) Ag-Jammin_ 1–2 (Biedler and Tu, 2003) RT1, RT2, Ag-R1 _1–11 (Biedler and Tu, 2003) R1, RT1 (Waldo) R1Bm (Xiong and Eickbush, 1988), SART1, TRAS1 (Anzai et al., 2001) Lian (Tu et al., 1998) Bilbo, Baggins1 (Kapitonov and Jurka, 2003b) LOA (Felger and Hunt, 1992) MosqI (Tu and Hill, 1999) Ag-I_ 1–7 (Biedler and Tu, 2003) I, I2, You JuanA (Mouches et al., 1992) Ag-Jockey_ 1–13, Ag-Jen _1–12, (Biedler and Tu, 2003) NCR1Cth1 (Blinov et al., 1997) JuanC (Agarwal et al., 1993) BS, Doc, F, G, Helena, Het-A, Jockey, Juan, TART, X AMY (BMC1, L1Bm; Abe et al., 1998) LDT1 (Garner and Slavicek, 1999) T1, Q, Ag-CR1 _1–29 (Biedler and Tu, 2003) DMCR1A (Kapitonov and Jurka, 2003b) Ag-Loner_ 1–3 (Biedler and Tu, 2003) Ag-Outcast_ 1–11 (Biedler and Tu, 2003) Sponge (Biedler and Tu, 2003) CM-gag (Bensaadi-Merchermek et al., 1997) Kurosawa, Kendo (Abe et al., 2002)

Diptera Lepidoptera Diptera Lepidoptera Diptera Diptera Diptera Diptera Diptera Diptera Lepidoptera

Anopheles gambiae Bombyx mori Drosophila melanogaster Bombyx mori Anopheles gambiae Anopheles gambiae Aedes aegypti Anopheles gambiae Anopheles gambiae Drosophila melanogaster Bombyx mori

Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Lepidoptera Lepidoptera Diptera Diptera Diptera Diptera Diptera Diptera Lepidoptera

Aedes aegypti Drosophila melanogaster Drosophila silvestris Aedes aegypti Anopheles gambiae Drosophila melanogaster Aedes aegypti Anopheles gambiae Chironomus thummi Culex pipiens Drosophila melanogaster Bombyx mori Lymantria dispar Anopheles gambiae Drosophila melanogaster Anopheles gambiae Anopheles gambiae Anopheles gambiae Culex pipiens Bombyx mandarina

Feilai (Tu, 1999) SINE200 (Holt et al., 2002) Cp1 (Liao et al., 1998)

Diptera Diptera Diptera

Aedes aegypti Anopheles gambiae Chironomus pallidivittatus

Continued

406 Insect Transposable Elements

Table 1 Continued Superfamily (clade)

Unclassified SINEs?

Element and referenceb

Order

Organismc,d

Twin (Feschotte et al., 2001) Bm1 (Adams et al., 1986) Lm1 (Bradfield et al., 1985) Maque (Tu, 2001b) DINE-1 (Locke et al., 1999) IE (Sun et al., 1991; Lampe and Willis, 1994)

Diptera Lepidoptera Orthoptera Diptera Diptera Lepidoptera

Culex pipiens Bombyx mori Locusta migratoria Anopheles gambiae Drosophila melanogaster Hyalophora cecropia

Diptera Diptera

Aedes aegypti Drosophila virilis

IV. Penelope-like retrotransposons Penelope

Penelope-like (Biedler and Tu, unpublished data) Penelope (Evgen’ev et al., 1997)

a

Classifications of LTR and non-LTR retrotransposons are according to Eickbush and Malik (2002). The citation is not always the original report of a TE. In some cases a review or an article describing the phylogenetic placement of the TE is cited instead. References for all D. melanogaster TEs are from Kaminker et al. (2002) unless otherwise noted. There are several PCR surveys of a few RNA-mediated TEs in different insects that are not listed in the table (Booth et al., 1994; Rongnoparut et al., 1998; Cook et al., 2000). c TEs in Drosophila species other than D. melanogaster are not listed unless they are the founding member of a group of TEs. d In some cases, TEs from only one species are listed although they were found in several closely related species. b

promoter and transcription termination sequences, the LTR retrotransposons have a flexible structure that allows gain, loss, and perhaps rearrangement of functional domains (Eickbush and Malik, 2002). While the RT and a few other protein domains perform the key function of making double-stranded cDNA, the acquisition of the IN activity allowed the integration of the cDNA in a way much like the mechanisms employed by DNA transposons. In fact, the IN domain and some of the prokaryotic and eukaryotic DNA transposases are believed to share a common origin (Capy et al., 1997; and see below). The acquisition of the env-like protein by some LTR retrotransposons such as gypsy confers the ability to leave the cell and become infectious retroviruses (Eickbush and Malik, 2002). 4.12.4.2.2. Non-LTR retrotransposons Twelve of the 17 clades of non-LTR retrotransposons have been found in insects (Eickbush and Malik, 2002; Biedler and Tu, 2003) (Table 1 and Figure 7). In fact, the founding members of many of these clades were discovered in insects. The characterization and classification of 104 families of non-LTR retrotransposons in A. gambiae have been recently reported (Biedler and Tu, 2003). The 104 A. gambiae families represent divergent lineages in eight previously established clades (R4, L1, RTE, R1, L2, CR1, Jockey, and I) and two new clades, Loner and Outcast. Representation appears to be biased toward the most derived clades in non-LTR retrotransposon evolution, especially the CR1 and Jockey clades, with 31 and 25 families respectively. All of the 31 A. gambiae CR1 elements are grouped together with

the Drosophila CR1 being a sister branch. On the other hand, there appear to be three groups (I, II, and III) of the A. gambiae Jockey elements, which may have different sister elements either from other mosquitoes or other dipterans (Biedler and Tu, 2003). 4.12.4.2.3. SINEs SINEs have not been extensively investigated in insects. Only a small number of them have been found as shown in Table 1. Insect SINEs characterized so far all belong to the tRNA-related group. The structural features of Feilai, a SINE discovered in Aedes aegypti, include a tRNA-related promoter region, a tRNA-unrelated conserved region, and a triplet tandem repeat at its 30 end (Figure 2). The Twin SINE family, which was discovered in Culex pipiens (Feschotte et al., 2001), consists of two tRNA-related regions separated by a 39 bp spacer. SINE200 from A. gambiae contains only one of the two conserved boxes found in tRNA-related Pol III promoters (Tu, unpublished data). DINE-1, a SINE from D. melanogaster, lacks the structural features of typical SINEs (Locke et al., 1999). Feilai, SINE200, and two other SINEs BM1 and Lm1 (Table 1) are all highly repetitive in their respective genomes. Twin is only moderately repetitive with a copy number of 500. Cp1, a Chironomus pallidivittatus SINE, inserts specifically to centromeric tandem repeats (Liao et al., 1998). 4.12.4.2.4. Two intriguing families: Maque and Penelope A family of very short interspersed repetitive elements named Maque has recently been found in A. gambiae. There are approximately 220

t0010

Table 2 DNA-mediated transposable elements in insects Superfamily/family

Defining features

I. Cut-and-paste transposons P 8 bp TSD, conserved transposase

hAT (hobo-Ac-Tam3)

8 bp TSD, conserved transposase

IS630-Tc1-mariner Tc1

TA TSD, DDE(D) catalytic triad DD34E

mariner d

DD34D

ITmD37D (maT)

DD37D

ITmD41D

DD41D

ITmD37E

DD37E

Pogoe

DDxD, long C-terminus

piggyBac

TTAA TSD, conserved transposase

Element and referencea

Order

Organismb,c

AgaP (8 subfamilies: Sarkar et al., 2003) P (Canonical, M, O, T), ProtoP (Hoppel), ProtoP_B (Kapitonov and Jurka, 2003b) Lu-P1, Lu-P2 (Perkins and Howells, 1992) P (Lee et al., 1999) hAT-type (Holt et al., 2002) hopper (Handler, 2003) Homer (Pinkerton et al., 1999) hobo (Calvi et al., 1991) Hermit (Coates et al., 1996) Hermes (Warren et al., 1994)

Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera

Anopheles gambiae Drosophila melanogaster Lucilia cuprina Musca domestica Anopheles gambiae Bactrocera dorsalis Bactrocera tryoni Drosophila melanogaster Lucilia cuprina Musca domestica

Quetzal (Ke et al., 1996) Crusoe (Hill et al., 2001), DD34E, Tiang, Topi, Tsessebe (Holt et al., 2002) Minos (Franz and Savakis, 1991) Bari1-2, HB, S, S2, Tc1 (Kaminker et al., 2002); Tc3-like (Shao et al., 2001) Unnamed element (Mikitani et al., 2000) mariner (Holt et al., 2002) D.mauritiana.mar1 (Robertson, 2002) mariner2 (Kaminker et al., 2002) A.mellifera.mar1 (Robertson, 2002) H.cecropia.mar1 (Robertson, 2002) ITmD37D (Holt et al., 2002) MdmaT1 (Claudianos et al., 2002) Bmmar1 (Robertson and Asplund, 1996), Bmmar6 (Robertson and Walden, 2003) Crmar2 (Gomulski et al., 2001) Tcp3.2 (Arends and Jehle, 2002; Robertson and Walden, 2003) A.aegypti.ITmD37E (Shao and Tu, 2001) A.gambiae.ITmD37E (Shao and Tu, 2001) Pogo-like (Holt et al., 2002) Pogo (Tudor et al., 1992) piggyBac (Holt et al., 2002; Sarkar et al., 2003) piggyBac (Handler and McCombs, 2000) looper1 (Kapitonov and Jurka, 2002) piggyBac (Sarkar et al., 2003) piggyBac (Cary et al., 1989)

Diptera Diptera Diptera Diptera Lepidoptera Diptera Diptera Diptera Hymenoptera Lepidoptera Diptera Diptera Lepidoptera Diptera Lepidoptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Lepidoptera Lepidoptera

Anopheles albimanus Anopheles gambiae Drosophila hydei Drosophila melanogaster Bombyx mori Anopheles gambiae Drosophila mauritiana Drosophila melanogaster Apis mellifera Hyalophora cecropia Anopheles gambiae Musca domestica Bombyx mori Ceratitis rosa Cydia pomonella Aedes aegypti Anopheles gambiae Anopheles gambiae Drosophila melanogaster Anopheles gambiae Bactrocera dorsalis Drosophila melanogaster Bombyx mori Trichoplusia ni

Continued

Table 2 Continued Superfamily/family

Defining features

Element and referencea

Order

Organismb,c

PIF-harbinger

3 bp TSD, conserved transposase 5 bp TSD, conserved transposase

PIF/harbinger-like (Biedler et al., unpublished data)

Diptera

Anopheles gambiae

Transib1_AG (Kapitonov and Jurka, 2003b)

Diptera

Anopheles gambiae

Transib1-4 (Kapitonov and Jurka, 2003b), Hopper (Bernstein et al., 1995) Ikirara (Romans et al., 1998) TECth1 (Wobus et al., 1990)

Diptera Diptera Diptera

Drosophila melanogaster Anopheles gambiae Chironomus thummi

Helitron1_AG, Helitron 2_AG (Kapitonov and Jurka, 2003b) Helitron (Kapitonov and Jurka, 2003b)

Diptera Diptera

Anopheles gambiae Drosophila melanogaster

Transib

Not classified II. Rolling circle transposons Helitron

III. MITEs mTATA TSD

No TSD, similarity to helicase DEC (Braquart et al., 1999)

m3bp (Tourist-like MITEs)

3 bp TSD

m4bp m7bp m8bp

4 bp (often TTAA) TSD 7 bp TSD 8 bp TSD

m9bp

9 bp TSD

Not classified

Cleoptera

Tenebrio molitor

Pony (Tu, 2000), Wujin (Tu, 1997); mTA_1–8 (Mao and Tu, unpublished data) TA-I-Ag, TA-II-Ag, TA-III-Ag, TA-IV-Ag, TA-V-Ag (Tu, 2001a) Mikado, Milord, Mimo, Mirza, Nemo (Feschotte et al., 2002) mPogo (Feschotte et al., 2002) m3bp_1–10 (Mao and Tu, unpublished data) Joey, TAA-I-Ag, TAA-II-Ag (Tu, 2001a), m3bp_1–19 (Biedler et al., unpublished data)

Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera Diptera

A MITE in a NOS gene (Luckhart and Rosenberg, 1999) Wukong, Wuneng (Tu, 1997), m4bp_1–6 (Mao and Tu, unpublished data) m7bp_1–3 (Mao and Tu, unpublished data) m8bp_1–10 (Mao and Tu, unpublished data) 8bp-I-Ag (Tu, 2001a), Pegasus (Besansky et al., 1996) Mar, Vege (Holyoake and Kidwell, 2003) m9bp_1–2 (Mao and Tu, unpublished data) Microuli (TTAA TSD, SIR: Tu and Orphanidis, 2001) MEC (5 bp TSD: Blinov et al., 1991) Mint (CA TSD, SIR: Feschotte et al., 2002) SGM-IS (SIR: Miller et al., 2000)

Aedes aegypti Anopheles gambiae Culex pipiens Drosophila melanogaster Aedes aegypti Anopheles gambiae Anopheles stephensi Aedes aegypti Aedes aegypti Aedes aegypti Anopheles gambiae Drosophila willistoni Aedes aegypti Aedes aegypti Chironomus thummi Culex pipiens Drosophila obscura

a The citation is not always the original report of a TE. In some cases a review or an article describing the phylogenetic placement of the TE is cited instead. Drosophila Foldback elements are not listed. There are several PCR surveys of a few DNA-mediated TEs in different insects that are not listed in the table (Daniels et al., 1990; Robertson, 1993; Bigot et al., 1994; Clark and Kidwell, 1997; Imwong et al., 2000; Green and Frommer, 2001). b TEs in Drosophila species other than D. melanogaster are not listed unless they are the founding member of a group of TEs. c In some cases, TEs from only one species are listed although they were found in several closely related species. d mariner sequences, full-length or fragment, have been found in a wide range of insects. The DD34D mariners consist of 6 subfamilies briggase, cecropia, elegans, irritans, mauritiana, and mellifera. See Shao and Tu (2001) for a reclassification of the IS630-Tc1-mariner superfamily. See Robertson (1993, 2002) for reviews on mariner evolution. e The status of Pogo is not certain. It is either considered a member of the IS630-Tc1-mariner superfamily or a separate superfamily (Shao and Tu, 2001; Robertson, 2002).

Insect Transposable Elements

copies of Maque. Only approximately 60 bp long, Maque has the appearance of a distinct transposition unit. The majority of Maque elements were flanked by 9–14 bp TSDs. Maque has several characteristics of non-LTR retrotransposons such as TSDs of variable length, imprecise 50 terminus, and CAA simple repeats at the 30 end. The evolutionary origin of Maque and the differences between Maque and other known retro-elements including SINEs is not yet known. The 50 end of Maque represents a strong stop position that caused frequent premature termination of reverse transcription is suggested (Tu, 2001b). Although no autonomous non-LTR retrotransposons have been found that share similar 30 sequences with Maque, there is a family of non-LTR retrotransposons, Ag-I-2 (Biedler and Tu, 2003) that has the same CAA tandem repeats at their 30 termini. It is possible that short sequences such as Maque that contain just the RT recognition signal could potentially contribute to the genesis of some primordial SINEs (Tu, 2001b). Penelope, another intriguing family, was discovered as a TE involved in the hybrid dysgenesis of crosses between field-collected and laboratory strains of D. virilis (Evgen’ev et al., 1997). It has a RT that is grouped with the RT from telomerase (Arkhipova et al., 2003). More strikingly, members of the Penelope family in bdelloid rotifers are able to retain their introns, which is inconsistent with a transposition mechanism involving an RNA intermediate. It was proposed that the Uri endonuclease domain found in all Penelope-like elements may allow them, at least in part, to use a DNA-mediated mechanism similar to that used by group I introns (Arkhipova et al., 2003). On the basis of these unique features, Penelope was classified as a unique group that is distinct from LTR and non-LTR retrotransposons. 4.12.4.3. DNA-Mediated TEs

4.12.4.3.1. Cut-and-paste DNA transposons The majority of DNA-mediated TEs are believed to transpose by a cut-and-paste mechanism. This group of elements is characterized by 10–200 bp TIRs flanking one or more ORFs that encode a transposase, the enzyme that performs the excision and integration (cut and paste) of the cognate TE DNA. Their copy number may be increased through a repair mechanism, or by transposing ahead of a replication fork (Finnegan, 1992; Craig, 2002). Different families/superfamilies of insect TEs in this group are listed in Table 2. Again, several insect TEs are the founding members of their respective families/superfamilies that have broad distributions. The families/superfamilies that have been found in

409

insects include IS630-Tc1-mariner, hAT, piggyBac, PIF/Harbinger, P, and Transib (Shao and Tu, 2001; Robertson, 2002; Kapitonov and Jurka, 2003b). Conserved transposase sequences and TSDs of specific sequence or length are the hallmarks of each family/superfamily. In vitro experiments have shown that, for a few cut-and-paste DNA transposons characterized so far, the transposase alone is sufficient to direct the transposition reactions by interacting with the TIRs and the insertion target (Plasterk et al., 1999). However, the transposition of P elements requires a host protein (inverted repeat binding protein, IRBP) that binds to its TIRs (Badge and Brookfield, 1997; Rio, 2002). This requirement may explain the lack of success in the early attempts to use P elements to transform insects other than Drosophila. The structural characteristics of representative elements from each family are shown in Figure 3 (see Chapter 4.13). A few TE families including PIF/Harbinger (Biedler et al., unpublished data) and Transib (Kapitonov and Jurka, 2003b), which were discovered in insects only recently, will be highlighted in this section. Recent expansion and reclassification of the IS630-Tc1-mariner superfamily will also be discussed (Shao and Tu, 2001). In a separate section on MITEs (see Section 4.12.4.3.2), two new groups of mosquito MITEs will be described that have 7 and 9 bp TSDs respectively (Mao and Tu, unpublished data). The discovery of MITEs with 9 bp TSDs provides a preliminary indication of the possible existence of Mutator-like transposons in insects, which have so far been only found in plants (Walbot and Rudenko, 2002). On the other hand, the 7 bp TSD MITEs could potentially lead to the discovery of an entirely new family of eukaryotic DNA transposons because 7 bp TSD elements have until now only been found in bacteria (Krebs et al., 1990; Mahillon and Chandler, 1998) (see Section 4.12.4.3.2.2). 4.12.4.3.1.1. Discovery of the ITmD37E transposon and the reclassification of the IS630-Tc1mariner (ITm) superfamily It was shown previously that some prokaryotic IS elements, eukaryotic Tc1 and mariner transposons, and eukaryotic retrotransposons and retroviruses form a megafamily which shares similar signature sequences or motifs in the catalytic domain of their respective transposase and IN (Capy et al., 1996, 1997). The common motif for this transposase–integrase megafamily is a conserved D(Asp)DE(Glu) or DDD catalytic triad. The distance between the first two Ds is variable while the distance between the last two residues in the catalytic triad is mostly invariable

410 Insect Transposable Elements

Insect Transposable Elements

for a given transposon family in eukaryotes, indicating functional importance. Within this megafamily, the eukaryotic DNA transposon families Tc1 and mariner and the bacterial IS630 element and its relatives in prokaryotes and ciliates comprise a superfamily, the IS630-Tc1-mariner superfamily, which is based on overall transposase similarities and a common TA dinucleotide insertion target (Henikoff, 1992; Doak et al., 1994; Robertson and Lampe, 1995; Capy et al., 1996; Shao and Tu, 2001). Tc1-like elements identified in fungi, invertebrates, and vertebrates all contain a DD34E motif while most mariner elements identified in flatworms, insects, and vertebrates contain a DD34D motif. A few TEs that contain DD37D and DD39D motifs were previously regarded as basal subfamilies, the max subfamily and mori subfamily respectively, of the mariner family (Robertson, 2002). A novel transposon, ITmD37E, was recently reported in a wide range of mosquito species (Shao and Tu, 2001). The ITmD37E transposases contain a conserved DD37E catalytic motif, which is unique among the reported transposons of the ITm superfamily. Sequence comparisons and phylogenetic analyses suggest that ITmD37E is a novel family (Figure 8). In addition, our phylogenetic analyses show that the mori subfamily (DD37D) and max subfamily (DD39D) of mariner may also be classified as two distinct families, namely the ITmD37D and ITmD39D families. In fact, ITmD37D (previously mori subfamily of mariner) is more closely related to Tc1 (DD34E) than other mariner elements (Figure 8). The recognition of the three new families ITmD37E, ITmD37D, and ITmD39D is consistent with the fact that they share family-specific catalytic motifs and similar TIRs. Claudianos and colleagues also noticed the need for reclassification of the DD37D transposons and named them the maT family (Claudianos et al., 2002). Finally, a group of transposons that contain a DD41D catalytic motif

411

have been found in the medfly Ceratitis rosa, establishing yet another family (Gomulski et al., 2001; Robertson and Walden, 2003), namely the ITmD41D family. In summary, according to recent analyses, the ITm superfamily can be organized in seven families including ITmD37E, ITmD37D, ITmD39D, ITmD41D, Tc1, mariner, pogo, and an unresolved clade which includes bacterial IS630-like elements and some fungal and ciliate transposons (Figure 8). pogo is an interesting case as it has a unique N-terminal DNA-binding domain and a long C-terminal domain rich in acidic residues, although it contains a DDxD catalytic domain related to the ITm transposons (Smit and Riggs, 1996). 4.12.4.3.1.2. Two new families: PIF/Harbinger and Transib PIF/Harbinger and Transib are two newly discovered families of DNA transposons in insects. PIF and related TEs Harbinger and Tc8 have previously been found in plants and nematodes (Kapitonov and Jurka, 1999; Le et al., 2001; Zhang et al., 2001) and they may have mobilized Tourist, one of the most abundant MITEs in plants (Zhang et al., 2001; Jiang et al., 2003). Analysis of the A. gambiae genome uncovered approximately 30 families of PIF/Harbinger-like transposons, although the majority of them are ‘‘fossil’’-like sequences (Biedler et al., unpublished data). Like all other PIF/Harbinger-like transposons, the A. gambiae PIF/Harbinger-like elements are characterized by 3 bp TSDs and transposase sequences similar to that of the bacterial IS5 elements (Zhang et al., 2001). A second new family, the Transib transposons, are so far found only in D. melanogster and A. gambiae (Kapitonov and Jurka, 2003b). They use a unique transposase and generate 5 bp TSDs. Transib transposons have resided in these two genomes for a long time as extensive in silico reconstruction had to be employed to trace back the ancestral full-length copies (Kapitonov and Jurka, 2003b).

Figure 7 Phylogenetic analysis of diverse non-LTR retrotransposons found in Anopheles gambiae. Phylogenetic analysis classifies 104 families of A. gambiae elements into two new clades and eight previously defined clades. The two new clades, Loner and Outcast, are in bold. Shown here is the neighbor-joining tree constructed using alignment of approximately 260 amino acids of the reverse transcriptase (RT) domain from non-LTR retrotransposons of A. gambiae and representative elements in different clades. The tree was rooted using RTs of three prokaryotic Group II introns (not shown). Maximum parsimony was also used which produced a similar phylogenetic tree (not shown). Confidence of the groupings was estimated using 500 bootstrap replications for both methods. The first and second numbers at a particular node represent the bootstrap values derived from 500 neighbor-joining and maximum parsimony analysis, respectively. Only the values for the major groupings (clades) that are above 50% are shown. The scale at bottom left indicates amino acid divergence. The names of elements from previously established clades are given but names of new A. gambiae non-LTR families are omitted to save space (see Biedler and Tu, 2003). Previously reported A. gambiae nonLTRs in the tree are Q, T1, RT1, and RT2 (Besansky, 1990; Besansky et al., 1992, 1994). (Reproduced with permission from Biedler, J., Tu, Z., 2003. Non-LTR retrotransposons in the African malaria mosquito, Anopheles gambiae: unprecedented diversity and evidence of recent activity. Mol. Biol. Evol. 20, 1811–1825; ß Oxford University Press.)

412 Insect Transposable Elements

Figure 8 Structural features and classification of the IS630-Tc1-mariner superfamily. (a) Structural features. The catalytic triad in the transposase is highlighted. The characteristic TA target site duplications (TSDs) flanking an IS630-Tc1-mariner are shown. The terminal inverted repeats (TIRs) specify the boundries of the element. Possible introns are not shown. (b) Phylogenetic relationship between members of the IS630-Tc1-mariner superfamily on the basis of the catalytic domain. The alignment used here was previously described (Shao and Tu, 2001). The tree shown here is an unrooted phylogram constructed using a minimum evolution algorithm. Two additional methods, neighbor-joining and maximum parsimony, were also used. Confidence of the groupings was estimated using 500 bootstrap replications. The bootstrap value represents percent of times that branches were grouped together at a particular node. The first, second, and third numbers represent the bootstrap value derived from minimum evolution, neighborjoining, and maximum parsimony analysis, respectively. Only the values for major groupings are shown. Various colors indicate different clades. All phylogenetic analyses were conducted using PAUP 4.0 b8 (Swofford, 2001). Note that a recently described group of transposons that contain a DD41D catalytic triad was not included here. They are a distinct group related to the DD37D transoposons (Gomulski et al., 2001). (Modified from Shao, H., Tu, Z., 2001. Expanding the diversity of the IS630-Tc1-mariner superfamily: discovery of a unique DD37E transposon and reclassification of the DD37D and DD39D transposons. Genetics 159, 1103–1115.)

4.12.4.3.2. MITEs As shown in Table 2, MITEs that share similar TSDs and TIRs with DNA transposons in the IS630-Tc1-mariner, hAT, piggyBac, PIF/Harbinger families have been found in three species of mosquitoes. A MITE that generates a specific TA TSD has also been reported in a coleopteran (Braquart et al., 1999). Two hAT-like MITEs have been recently found in D. willistoni (Holyoake and Kidwell, 2003) and a deletion-derivative of the pogo transposon has been found in D. melanogaster (Feschotte et al., 2002).

The relationship between PIF/Harbinger-like DNA transposons and related Tourist-like MITEs in A. gambiae is interesting (Biedler et al., unpublished data). There are multiple PIF/Harbinger-like DNA transposon families in A. gambiae, all of which have the characteristic AT-rich 3 bp TSDs. Although the TIRs are highly conserved between different copies of a family, they are often variable between different PIF/Harbinger-like transposon families. Only two MITE families were found to be apparent deletion derivatives of two PIF/Harbinger-like

Insect Transposable Elements

transposons (Figure 4). On the other hand, 20 families of the Tourist-like MITEs were found to share similar TSDs and TIRs with their respective PIF/Harbinger-like transposons although no similarities were found between the internal sequences of these MITEs and any of the PIF/Harbinger-like transposons. The implication of these results on the evolution of MITEs is discussed elsewhere (see Sections 4.12.2 and 4.12.6). 4.12.4.3.2.1. m9bp, MITEs with 9 bp TSDs Two novel groups of MITEs have been discovered in Aedes aegypti. One group, m9bp, includes two families of putative MITEs that have 9 bp TSDs (Mao and Tu, unpublished data). These two families are moderately repetitive and evidence of insertion has been found for both families. Several plant MITEs have 9 bp TSDs (Charrier et al., 1999; Feschotte et al., 2002) and they may be mobilized by Mutator-like transposons only found in plants (Walbot and Rudenko, 2002). The discovery of 9 bp TSD MITEs in Aedes aegypti is the first in animals, which may indicate a broader distribution of the Mutator-like transposons.

p0200

p0205

4.12.4.3.2.2. m7bp, MITEs with 7 bp TSDs Three families of putative MITEs that have 7 bp TSDs (m7bp_1, m7bp_2, m7bp_3) have been found in Aedes aegypti (Mao and Tu, unpublished data). These families are moderately repetitive and evidence of insertion has been found for all three families. These 7 bp TSD MITEs could potentially lead to the discovery of an entirely new family of eukaryotic DNA transposons, as 7 bp TSD elements have until now only been found in bacteria (Krebs et al., 1990; Mahillon and Chandler, 1998). No full-length protein-encoding DNA transposons that produce 7 bp TSDs have been identified in A. aegypti yet. However, this is within expectation considering the fragmented nature of the BAC-end sequences in the current Aedes aegypti database. 4.12.4.3.2.3. Microuli, a miniature subterminal inverted-repeat TE Microuli is a family of small (200 bp) and highly AT rich (68.8–72.6%) TEs found in Aedes aegypti that do not have any coding capacity (Tu and Orphanidis, 2001). There is a 61 to 62 bp internal subterminal inverted repeat as well as a 7 bp subterminal inverted repeat 11 bp from the two termini. In addition, there are three imperfect subterminal direct repeats near the 50 end. All of the above characteristics clearly resemble the structural features of MITEs. The only feature that separates Microuli from MITEs is that Microuli elements lack TIRs. Therefore, we use the phrase ‘‘miniature

413

subterminal inverted-repeat transposable elements,’’ or MSITEs, to refer to the structural characteristics of the Microuli elements. Short insertion sequences that contain subterminal inverted repeats but lack TIRs have been identified in the genomes of rice and a Culex mosquito (Song et al., 1998; Feschotte and Mouches, 2000a). Fourteen of the 19 nucleotides at the 50 (and only 50 ) terminus of Microuli are identical to the TIR of Wuneng, a previously characterized MITE in Aedes aegypti (Tu, 1997). Both Microuli and Wuneng insert specifically into the TTAA target. It has been suggested that MITEs and the autonomous DNA transposons share the same transposition machinery based on common TIRs (Feschotte et al., 2002). Then how did Microuli transpose without the TIRs? The three subterminal direct repeats could potentially be the binding sites for transposases because subterminal inverted repeats and subterminal direct repeats have been shown to bind transposases in several autonomous DNA transposons (Morgan and Middleton, 1990; Beall and Rio, 1997; Becker and Kunze, 1997). It remains unclear how the termini of Microuli are determined at the strand cleavage step without the TIR. The TTAA target duplication plus a 3 bp TIR are essential for the excision of the autonomous transposon piggyBac (Bauser et al., 1999). Therefore, it is possible that Microuli may also be able to use the TTAA target sequence as part of the signal for recombination. It is tempting to hypothesize that some MITEs could evolve from MSITEs through mutation and/or recombination events at the termini which would result in TIRs. 4.12.4.3.3. Helitrons Helitrons have been found in D. melanogaster and Anopheles gambiae, which use a rolling-circle mechanism of transposition (Kapitonov and Jurka, 2001; Kapitonov and Jurka, 2003b) (Table 2). Insect Helitrons has several characteristics including short specific terminal sequences (50 TC and 30 CTAG), a 30 hairpin, and the lack of TSDs. Instead of a cut-and-paste transposase, Helitron1 in A. gambiae encodes an intronless protein including domains similar to helicase and replication initiation protein. There are approximately 100 copies of Helitron elements in A. gambiae that form 10 distinct families (Kapitonov and Jurka, 2003b). In summary, the diversity demonstrated by insect TEs encompasses all classes and types of eukaryotic elements. Analyses of insect TEs have more than once led to discoveries that broadly impacted the field of TE research. Because of extensive genetic studies, D. melanogaster has long served as the launching pad for the discovery of novel families

414 Insect Transposable Elements

of TEs in eukaryotes. The advent of genomics has provided the opportunity for the discovery of novel TEs in a wide range of organisms including nondrosophilid insects. As demonstrated above, greater TE diversity may be revealed and new insights may be gained from these genomic analyses.

4.12.5. Search for Active TEs in Insects Active TEs may be used as tools for genetic manipulation of insects in basic and applied research (see Section 4.12.9). In addition, the behavior of TEs in host genomes and their spread in natural populations may be studied by monitoring active TE families. It is therefore highly desirable to isolate active copies of TEs. As described above (see Section 4.12.3), TEs discovered from observations of genetic mutations tend to result from active transposition events. Although several active TEs were discovered in this manner, this discovery process relies heavily on fortuitous events. Several methods that, when used in concert, could provide a systematic approach to uncover active TEs in insect genomes are described below. 4.12.5.1. Identification of Potentially Active TEs on the Basis of Bioinformatics Analysis

As discussed above, the ongoing genome revolution has produced an immense amount of sequence data from which diverse TEs can be identified in various insect genomes. The computational programs described above (see Section 4.12.3) can greatly facilitate the discovery and characterization of a large number of TE families. Unfortunately, the vast majority of TEs have accumulated inactivating mutations during evolution, rendering the discovery of active TEs the difficult task of ‘‘finding needles in a haystack.’’ Bioinformatics analysis can provide leads to potentially active candidates that can be studied further. For example, using a semiautomated reiterative search strategy, many potentially active families of non-LTR retrotransposons in the A. gambiae genome were identified (Biedler and Tu, 2003). Here candidate families were identified based on sequence characteristics, which include the presence of full-length elements, intact ORFs, multiple copies with high nucleotide identity, and the presence of TSDs. High nucleotide identity indicates recent amplification from a source element, without enough time for divergence caused by nucleotide substitution and other mutations. It should be emphasized that sequence analysis can only provide leads for further analysis. For example, high sequence identity between copies of a TE family may not always indicate recent transposition activity

because it can also result from gene conversion events. 4.12.5.2. Detection of TE Transcription

Transcription is a required step during transposition of the RNA-mediated TEs. Although DNA-mediated TEs do not use RNA as an intermediate, transcription is required for production of transposase proteins. Therefore, the detection of transcription may offer further support for an active family in both classes of TEs. Transcription can be inferred if a match was found in an expressed sequence tag (EST) database to a TE sequence from the same organism. For example, 21 families of non-LTR retrotransposons had significant hits when BLAST searches were carried out against over 94 000 A. gambiae ESTs downloaded from NCBI (Biedler and Tu, 2003). Transcription of TEs can also be detected experimentally by real time polymerase chain reaction (RT-PCR), Northern blot, and even microarray. The source of mRNA may affect the outcome of these experiments because the activity of some TEs may be temporally and spatially controlled. It has been shown that TE activity can be elevated during the culturing of mammalian and plant cells (Wessler, 1996; Grandbastien, 1998; Liu and Wendel, 2000; Kazazian and Goodier, 2002). Different cell lines are available for a number of insect species. One caveat of the above approach is that, transcripts shown by either experimental detection or EST analysis could arise from spurious transcription. These transcripts could originate by transcription from a nearby host promoter. 4.12.5.3. Detection of Transposition Events by TE Display

TE display (Van den Broeck et al., 1998; Casa et al., 2000; Biedler et al., 2003) is a sensentive and reproducible experimental method to detect TE insertions. We have used it recently to study insertion site polymorphisms of endogenous insect TEs (Biedler et al., 2003) (see Sections 4.12.7 and 4.12.9). It is a modified form of amplified fragment length polymorphism (AFLP), the difference being that one primer is designed according to a TE sequence (Figure 9). First, genomic DNA is digested using a four-base restriction enzyme such as BfaI and then ligated to an adapter sequence. This is followed by two rounds of PCR. During the second PCR, a radioactive-labeled nested primer specific for the TE sequence is used following a touch-down protocol. After the second PCR, products are run on a sequencing gel and visualized by autoradiography. TE display is a powerful tool for genome-wide analysis of TE insertions and for detection of new insertions due to transposition (De Keukeleire et al.,

Insect Transposable Elements

415

Figure 9 TE display, a method to scan multiple insertion sites of a TE in the genome. (a) Principle of TE display, which is a modified form of amplified fragment length polymorphism (AFLP). The difference is that TE-specific primers (F1 and F2) are used in addition to the adaptor primer (R1). F2 is labeled as shown by the asterisk. (b) Partial image of a TE display using primers for the Pegasus element with eight female individuals from an Anopheles gambiae colony (GAMCAM) originally collected from Cameroon (Biedler et al., 2003). The eight samples on the left are amplified with a Pegasus-specific primer Peg-F2. The eight samples on the right are the same as those on the left except they were amplified with primer Peg-F3, which is designed to amplify a product smaller by three bases. The three base shift is clearly observable. A 10 bp ladder is shown on the right. Bands from a TE display gel were reamplified and sequenced, showing that they contained Pegasus sequences as well as flanking genomic and adapter sequences in the expected order (not shown). Comigrating bands among different individuals had the same flanking genomic sequence, indicating that they were from the same genomic locus. TE displays have been also developed using two highly reiterated SINEs, SINE200 in A. gambiae and Feilai in Aedes aegypti (not shown). (Modified from Biedler, J., Qi, Y., Holligan, D., Della Torre, A., Wessler, S., et al., 2003. Transposable element (TE) display and rapid detection of TE insertion polymorphism in the Anopheles gambiae species complex. Insect Mol. Biol. 12, 211–216.)

2001). It offers a higher degree of sensitivity and resolution than genomic Southern blot analysis. TE display has been used to detect somatic cell transposition (De Keukeleire et al., 2001), simply by looking for the presence of new bands that represent newly transposed copies of a TE. A caveat of this approach is that a change in a restriction site may also result in new TE display bands. The same method may also be used to identify germline transposition by comparing TE display patterns of parent insects with the patterns of a large number of offspring. Alternatively, one could take advantage

of the possibility that some TEs are activated in cell culture. Using TE display, one may be able to identify active families by comparing the relative abundance of a TE in cultured cells with that in individuals from different strains of the same species (Jiang et al., 2003). This approach is based on the assumption that some TE families may be more active in cultured cells than in live organisms. After TE display, one side of the new insertional copy and its associated flanking sequence can be recovered because the band can be reamplified and sequenced. To recover the entire sequence of the

416 Insect Transposable Elements

newly inserted copy, one has to rely on mapping the insertion site to the genome if the genome sequence is available. Otherwise, further PCR or screening of a genomic library is necessary to recover the entire copy and both sides of the flanking sequence. The identified insertional copy itself may or may not be active because it could have been mobilized by a trans-acting protein that is encoded by an active TE in the genome. However, the experiment described above can demonstrate in vivo or ex vivo transposition events and the recovered insertional copy may lead to the identification of the autonomous active TE on the basis of shared cis-acting sequences such as TIRs. 4.12.5.4. Detection of Transposition Events by Inverse PCR

Actively transposing DNA-mediated TEs can be identified as extrachromsomal DNA in the form of linear or circular intermediates or byproducts (Arca et al., 1997; Gorbunova and Levy, 1997) (see Chapter 4.13). Using a set of outward-orienting primers within the TE, the circular extrachromosomal copies may be amplified, which could serve as evidence of active excision or transposition. However, head-to-head copies of the same TE in the genome could also produce PCR products when outwardorienting primers are used, which must be ruled out by sequencing and further analysis. Extrachromosomal circles of Hermes (see Chapter 4.13) and Pegasus (Coy and Tu, unpublished data) have been identified. Imprecise excision had occurred when Pegasus is excised from its genomic location within the A. gambiae genome. 4.12.5.5. Transposition Assay, Reconstruction, and Genetic Screen

Transposition assays can be used to directly assess the functionalities of both the cis-(TIRs) and the trans- (transposase) components of a DNA transposon, allowing the demonstration of autonomous transposition events (see Chapter 4.13). In addition, transposition assays have also been established for the detection of retrotransposition of non-LTR retrotransposons (Jensen et al., 1994; Ostertag et al., 2000). Recently, a molecular reconstruction approach has been developed to restore inactivated copies of a vertebrate transposon, Sleeping Beauty (Ivics et al., 1997), but such an approach requires extensive phylogenetic analysis and elaborate reconstructions (Ivics et al., 1997). As an alternative, a genetic screen based on a bacterial system has been developed to identify hyperactive copies of an insect mariner transposon among randomly mutated copies (Lampe et al., 1999). This approach can potentially be used to screen for active copies of

transposons that do not require specific host factors. In summary, the progress in insect genome projects and the development and application of the methods described in this section will greatly facilitate searches for active TEs. The task of ‘‘finding needles in a haystack’’ could potentially be replaced by targeted and more efficient investigations.

4.12.6. Evolution of Insect TEs The evolutionary dynamics of TEs are complex, which is in part due to their replication and their interactions with the host genome. The intricate dynamics between TEs and their host genomes are further complicated by the ability of some TEs to cross species barriers and spread in a new genome by horizontal transfer. Horizontal transfer may be an important part of the life cycle of some TEs and contribute to their continued success during evolution (Silva et al., 2004). While the broad distribution of both RNA-mediated TEs and DNA-mediated TEs in all eukaryotic groups is evidence of the long-term evolutionary success of TEs, the two TE classes may have adopted different strategies, for which several insect TEs in both classes provide good examples. 4.12.6.1. Genomic Considerations of TE Evolution

It has been hypothesized that TE insertions may present three types of potentially deleterious effects including: (1) insertional mutagenesis which may disrupt gene function and/or regulation; (2) transcriptional/translational cost of the production of TE transcripts and proteins; and (3) ectopic recombination between homologous copies of TEs in different chromosomal locations that may result in duplication, deletion, and a new linkage relationship between genes (Nuzhdin, 1999; Bartolome et al., 2002; Kidwell and Lisch, 2002; Rizzon et al., 2002; Petrov et al., 2003). The costs of having TEs may also include the cost associated with DNA replication when TEs occupy a large fraction of the genome. Obviously, these hypotheses are not mutually exclusive. This section discusses the intragenomic dynamics of TE–host interaction. The population dynamics affecting the spread of TEs in insects, which is also important for TE evolution, will be discussed below (see Section 4.12.7). 4.12.6.1.1. Self-regulation of insect TEs Both TEdriven mechanisms (self-regulation) and host-driven mechanisms (host control) have been shown to affect TE activities in insects. Self-regulation has been shown for mariner and P elements in Drosophila

Insect Transposable Elements

(Hartl et al., 1997; Kidwell and Lisch, 2001). In the case of the Drosophila P element, self-regulation is achieved through the activities of at least two types of element-encoded repressors. In the case of mariner, several mechanisms may be involved including overproduction inhibition (an increase in the amount of transposase results in a decrease in net transposase activity), missense mutation effects (defective transposase encoded by missense copies interfering with functional transposase), and titration effects by inactive copies. In this regard, it is interesting to note that several hyperactive mutants of an active mariner, originally discovered in the horn fly, have been isolated (Lampe et al., 1999). This suggests that the horn fly mariner has not evolved for maximal activity. 4.12.6.1.2. Host control of TEs in insects Host control of TE activity can either be targeted at a particular family of TEs or a large group of TEs in general. Family-specific host control has been demonstrated in several cases (Labrador and Corces, 2002). The best example so far is the control of gypsy activity by a genetic locus flamenco in D. melanogaster (Bucheton, 1995). Homozygous female mutants of flamenco, which is an X-linked recessive gene that represses the transposition of gypsy, produce progeny that show high rates of gypsy transposition. The relief of the suppression of gypsy activity in flamenco mutants, which are also called permissive mutants, functions through maternal factors. Another example of host control is the P element which only transposes in the germline because correct splicing of the P transcript only occurs in germ cells. The tissue-specificity of the P element allows it to transmit efficiently to the next generation while minimizing potential damage to the host. Two host genes have been implicated in the inhibition of P activity in somatic cells (Siebel et al., 1992, 1995). RNA interference (RNAi), a mechanism that confers posttranscriptional degradation of mRNA on the basis of homology to small fragments of double-stranded RNA, has been implicated as a host defense mechanism against a broad spectrum of TEs in the nematode Caenorhabditis elegans (Ketting et al., 1999; Tabara et al., 1999). RNAi has been shown to function in Drosophila and a few other insects as well (Misquitta et al., 1999; Hammond et al., 2000). The potential role of RNAi as a general control method against TE activity in Drosophila has recently been proposed on the basis of cosuppression of the I element by an increasing number of I-related transgenes (Jensen et al., 1999; Labrador and Corces, 2002). It has been proposed that RNAi has evolved as a host

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defense mechanism against the invasion by TEs and viruses (review: Fedoroff, 2002). In addition to posttranscriptional degradation, this defense system may also be involved in the formation of transcriptionally inactive heterochromatin where TEs and other repeats concentrate (Couzin, 2002). 4.12.6.1.3. Nonrandom distribution of insect TEs Patterns of nonrandom TE distribution have been shown in both D. melanogaster and A. gambiae (Bartolome et al., 2002; Holt et al., 2002; Kapitonov and Jurka, 2003b). TEs tend to accumulate in heterochromatin. Such a distribution bias could result either from preferential TE insertion, or selection against insertions in euchromatic regions, or both. Bartolome and colleagues suggested that the abundance of TEs is more strongly associated with local recombination rates (Bartolome et al., 2002), which are low in heterochromatic regions, rather than with gene density. They argue that this association is consistent with the hypothesis that selection against harmful effects of ectopic recombination is a major force opposing TE spread. However, selection against insertional mutagenesis is also at work as shown by the absence of insertions in coding regions. The insertional bias of P elements has been demonstrated recently during genome-scale P mutagenesis analysis (Spradling et al., 1995, 1999). Therefore insertion bias may contribute to the biased pattern of TE distribution in insects. A related topic here is the suggestion that concentrations of TE insertions in the Drosophila Y chromosome may have contributed to the evolutionary process leading to its inactivation (Labrador and Corces, 2002). It should be noted that not all TEs have a bias towards heterochromatic or recombination-deprived regions. On the basis of analysis of limited gene sequences, it was shown that Aedes aegypti MITEs tend to be associated with the noncoding regions within or near genes (Tu, 1997), which is similar to what has been observed for plant MITEs (Zhang et al., 2000). 4.12.6.1.4. Autonomous and nonautonomous TEs In addition to the genomic interactions described above (see Section 4.12.6.1.3), most TEs have to contend with the fact that defective copies are often generated during or after transposition. This process could contribute to self-regulation as discussed above (see Section 4.12.6.1.1). It can also lead to total inactivation and ultimate extinction of a TE as the inactive TE population eventually overwhelms the active copies (Eickbush and Malik, 2002). Therefore, the replicative ability that is responsible for the success of the TE may also lead

p0290

418 Insect Transposable Elements

to its inactivation in a genome. Interestingly, some nonautonomous TEs have been very successful with regard to amplification, although the mechanisms that contribute to their success are not entirely clear. For example, SINEs found in insect genomes including Aedes aegypti, Anopheles gambiae, and Bombyx mori (Table 1) all contain thousands of copies. Similarly, most of the MITEs found in insects are also highly reiterated although low-copy-number families are also found (Tu, 1997, 2000, 2001a). The small size of MITEs and SINEs may confer less deleterious effects on the host, either because they are less efficient substrates for homologous recombination (Petrov et al., 2003) or because their impact on neighboring genes may be less severe. Therefore reduced selection pressure as well as other properties inherent to MITEs and SINEs could contribute to their apparent success. It is a fascinating question as to how SINEs and MITEs affect the evolution of the autonomous TEs that mobilize them. 4.12.6.2. Vertical Transmission and Horizontal Transfer of Insect TEs

As described above (see Section 4.12.6.1), the replicative ability that is responsible for the success of a TE may in some cases lead to its inactivation by generating defective copies or by activating host

control mechanisms. This is especially true for DNA transposons. Therefore, the ability to escape the above vertical inactivation by invading a new genome would greatly enhance the evolutionary success of DNA transposons. However, not all TEs have adopted this life cycle of invasion, amplification, senescence, and new invasion (Hartl et al., 1997; Eickbush and Malik, 2002; Robertson, 2002) (Figure 10). 4.12.6.2.1. Detection of horizontal transfer The occurrence of horizontal transfer can be supported in various degrees by three types of evidence (Silva et al., 2004). First, detection of elements with a high level of sequence similarity in divergent taxa will offer strong support for horizontal transfer although variable rates of sequence change should be considered. Second, detection of phylogenetic incongruence between TEs and their hosts will also provide relatively strong support for horizontal transfer. However, this alone will not be convincing, especially in light of the high levels of intragenomic diversity of TE families observed in insects. In other words, the existence of multiple TE lineages could confound the phylogenetic analysis as paralogous lineages may be treated as orthologous ones. Finally, horizontal transfer may be inferred when ‘‘patchy’’ distribution of a TE among closely related taxa is observed. This type of support is weak as loss of

Figure 10 A model of the evolutionary dynamics of TEs in eukaryotic genomes. This hypothetical model incorporates recent work by several groups (Hartl et al., 1997; Lampe et al., 2001; Silva et al., 2004). Some aspects of this model are better suited for DNA transposons that generally have a high propensity for horizontal transfer. Three possible alternatives to inactivation and stochostic loss are highlighted (A, B, and C).

Insect Transposable Elements

TEs from sister taxa may result from a phenomenon similar to assortment of an ancestral polymorphism (Silva et al., 2004). 4.12.6.2.2. Horizontal transfer and vertical transmission in insects: differences between different groups of TEs The first case of eukaryotic horizontal transfer was reported in Drosophila, where the P element was shown to have invaded the D. melanogaster genome during the last century from a species in the D. willistoni group (review: Kidwell, 1992). Evidence for this lateral event includes all three types of support described above and is therefore widely accepted (Silva et al., 2004). Further analyses of a large number of P element sequences from a number of Drosophila species showed that many horizontal transfer events must have occurred to account for the current distribution pattern of P in Drosophila (Silva and Kidwell, 2000; Silva et al., 2004). Another spectacular case of horizontal transfer, which was also initially discovered in insects, involves the mariner transposons. The mariner transposon family has been implicated in hundreds or more horizontal transfer events among a wide range of animal species including a large number of insects across different orders (Robertson, 1993, 2002). It has been shown that the newly discovered ITmD37E transposons have also been involved in relatively frequent horizontal transfer events among different mosquito species (Shao and Tu, 2001, unpublished data). Finally, horizontal transfers of hobo and piggyBac has been shown in flies and moths (Bonnivard et al., 2000; Handler and McCombs, 2000). It has been proposed that horizontal transfer may be a key link in the life cycle of those DNA TEs that are not intricately associated with their host biology, allowing them to escape the vertical inactivation and stochastic loss (Hartl et al., 1997). Thus, these types of TEs may be regarded as ‘‘resident aliens’’ in a host (Plasterk et al., 1999). It has been shown that selection of some mariner transposases acts only during horizontal transfer, which is consistent with the invasion–inactivation–escape model (Lampe et al., 2003). Horizontal transfer has also been shown for LTR retrotransposons in insects. A clear example involves the copia element in Drosophila (Jordan et al., 1999). The copia elements in D. melanogaster and D. willistoni, two divergent species that separated more than 40 million years ago, showed less than 1% nucleotide difference. It appears that copia jumped from D. melanogaster to D. willistoni. Horizontal transfer of the Drosophila gypsy

419

element has also been reported (Terzian et al., 2000; Vazquez-Manrique et al., 2000). Possible horizontal transfer of the Drosophila nonLTR retrotransposon Jockey was suggested on the basis of phylogenetic incongruence (Mizrokhi and Mazo, 1990). However, Malik and colleagues recently showed that an analysis that takes into account the rates of evolution was consistent with vertical transmission of the Jockey elements in Drosophila (Malik et al., 1999). Eickbush and Malik further suggest that no convincing evidence exists for the horizontal transfer of any non-LTR retrotransposon during the past 600 million years (Eickbush and Malik, 2002). In addition, strict vertical transmission of two non-LTR retrotransposons R1 and R2 have been shown in Drosophila (Eickbush and Eickbush, 1995). In these cases, the TE phylogenies well reflect those of the host species. Furthermore, it was shown that the relationship between R2 elements from several insect orders including Diptera, Lepidoptera, Coleoptera, and Hymenoptera is consistent with vertical transmission (Eickbush, 2002). 4.12.6.2.3. Possible reasons for differing propensities for horizontal transfer Two reasons have been proposed to explain the apparent differences in the prevalence of horizontal transfer events between DNA transposons and non-LTR retrotransposons (Eickbush and Malik, 2002). The first is that DNA transposons need horizontal transfer for their long-term survival but non-LTR retrotransposons appear not to be dependent on such rare evolutionary events. Defective copies of DNA transposons retain the ability to be transposed as long as they have the cis-acting signals such as the TIRs. This indiscrimination leads to the inevitable fate of inactivation of the entire transposon family. Therefore, horizontal transfer offers a much-needed escape from the above vertical inactivation, which greatly enhances the evolutionary success of DNA transposons. On the other hand, it has been shown that the RT of non-LTR retrotransposons tends to associate with the mRNA molecules from which they were translated (Wei et al., 2001). This cispreference would bias transposition events in favor of the active elements, thus providing a mechanism to sustain the non-LTR retrotransposons. However, the cis-preference is not enough to prevent the highly successful retrotransposition of SINEs in insects (Adams et al., 1986; Tu, 1999) and other organisms (Lander et al., 2001), which presumably borrows the retrotransposition machinery from non-LTR retrotransposons. It will be interesting to see how SINEs affect the evolution of their

420 Insect Transposable Elements

non-LTR retrotransposon ‘‘partners.’’ The second explanation is that DNA transposons may be more predisposed to horizontal transfer than non-LTR retrotransposons. The transposition process of DNA transposons involves an extrachromosomal DNA intermediate, which may facilitate horizontal transfer (Eickbush and Malik, 2002) (see Section 4.12.6.2.4). DNA transposons use their transposase for integration, which may be less dependent on host repair machinery than non-LTR retrotransposons and thus not as restricted to its original host. LTR retrotransposons form an extrachromosomal DNA intermediate and use transposase-like IN for integration. Therefore, LTR retrotransposons have access to the same horizontal transfer mechanisms as the DNA transposons although their life cycle may not require horizontal transfer because defective copies are not thought to be a major factor (Eickbush and Malik, 2002). It should be noted that the above are general statements and that the propensity for horizontal transfer may vary among individual families within the three groups discussed here.

showed a large number of lineages of non-LTR retrotransposons of the CR1 and Jockey clades in A. gambiae (Biedler and Tu, 2003) (Figure 7). Given the presence of multiple recently active lineages within the CR1 and Jockey clades, it is tempting to speculate that the observed diversity may be driven by competition among different non-LTR families or by attempts to escape suppressive mechanisms imposed by the host. On the other hand, some TEs are recruited for host functions and thus become ‘‘domesticated’’ (Lander et al., 2001; Kidwell and Lisch, 2002). This type of molecular domestication is the ultimate case of trading ‘‘freedom’’ for ‘‘security.’’ It allows TEs to sustain and positively impact the host, examples of which will be discussed below (see Section 4.12.8). Strictly speaking, these domesticated TEs are no longer TEs. However, it is theoretically possible that these ‘‘domesticated’’ TEs could revert back to their ‘‘old ways’’ on rare occasions.

4.12.6.2.4. Mechanisms of horizontal transfer Mechanisms of horizontal transfer are poorly understood although direct transfer of the extrachromosomal DNA intermediate and indirect transfer through a viral vector have been proposed as possible mechanisms (Eickbush and Malik, 2002; Silva et al., 2004). Geographical and temporal overlap between the donor and recipient host species may be essential. An intriguing case of horizontal transfer of a mariner element between a parasitoid wasp and its lepidopteran host offers a good example of such overlap (Yoshiyama et al., 2001).

High levels of TE diversity have been reported in the two insect genomes in which large-scale systematic analyses have been carried out (Holt et al., 2002; Kaminker et al., 2002; Kapitonov and Jurka, 2003b; Biedler and Tu, 2003) (Figures 4 and 7). The evolutionary process that generated this diversity may also be quite diverse. It is possible that the evolution of some TEs may be a complex mix of both vertical transmission and horizontal transfer events. Parsing out the results of intragenomic diversification from those of horizontal transfer events may require additional data from related species. Understanding the process responsible for the intragenomic diversity of insect TEs and the potential interactions between different TE families in insect genomes will be both challenging and rewarding. A summary of the current hypothesis on TE evolution is illustrated in Figure 10. Some aspects of this model are better suited for DNA transposons that have a high propensity for horizontal transfer.

4.12.6.3. Other Possible Evolutionary Strategies

In addition to horizontal transfer and vertical extinction, recent studies suggest that there might be a third way, or an alternative strategy, which may be adopted by some TEs (Lampe et al., 2001). On the basis of the loss of interaction between mariner transposons of slightly changed TIRs, it was proposed that intraspecific or intragenomic diversification of mariner transposons may allow the newly diverged mariner to start a new lineage. Although this requires the coevolutionary events to occur in both the transposase and the TIRs, this scenario would provide the transposon the opportunity to escape vertical inactivation because it is now virtually a brand new element in a virgin genome because of the loss of interaction between itself and its relatives in the genome. Genome sequencing has provided increasing opportunity to survey the diversity of different families of TEs. Our recent analysis

4.12.6.4. Understanding the Intragenomic Diversity of Insect TEs

4.12.7. TEs in Insect Populations 4.12.7.1. Fundamental Questions and Practical Relevance

In general, the increase of TE copy number through transposition is balanced by selective forces against the potential genetic load of TEs on host fitness (Nuzhdin, 1999). The control of transposition rate of TEs and other mechanisms to minimize their deleterious effects has been discussed (see Section 4.12.6). The population dynamics affecting the

Insect Transposable Elements

spread of TEs in insect genomes is described. Earlier work on TEs in Drosophila populations suggest that the copy numbers in euchromatic regions are low and most euchromatic copies exist at very low frequency (300 50–150 7–10 125 5–12 1–8 15 53–60 172 0.37 0.8 6–10 5.9 1.5–3.0

n.r. n.r. Shake flask T-flask 10 l bioreactor Shake flask 5 l reactor Spinner flask 30 l bioreactor 6-well plate 6-well plate 6-well plate Shake flask Spinner flask 3 l reactor 15 cm dish T-flask Shake flask n.r. Roller bottle 1 l reactor 23 l reactor 1 l suspension Roller bottles Spinner flasks Shake flasks Shake flask 8 l bioreactors n.r. 20 ml cultures 60 l bioreactor 4 l bioreactor Spinner flask Shake flask T-flask 1 l bioreactor T-flask Spinner flask Roller bottle suspension 2.5 l bioreactor

Steiner et al. (1988) Davis et al. (1992) DiPersio et al. (1992) Kang et al. (1992) Gierse et al. (1993) Tomlinson et al. (1993) Murphy et al. (1993) Lowe (1994) Kurkela et al. (1995) Bonning and Hammock (1995) Bonning and Hammock (1995) Bonning and Hammock (1995) Bonning and Hammock (1995) Bonning and Hammock (1995) Brown et al. (1995) Sugiura et al. (1995) Ciaccia et al. (1995) Homa et al. (1995) Brown et al. (1995) Geisse et al. (1996) Sorci-Thomas et al. (1996) Mathews et al. (1996) Withers et al. (1996) Winzerling et al. (1996) Mathews et al. (1996) Thordarson et al. (1996) Airenne et al. (1997) George et al. (1997) Wester et al. (1997) McQueney et al. (1998) Zhang et al. (1998) Canaan et al. (1998) Sviridov et al. (1999) Willard et al. (2000) Das et al. (2000) Pereira et al. (2001) Wang et al. (2001) Sadatmansoori et al. (2001) Ragunath et al. (2002) Ji et al. (2003) Ding et al. (2003)

AcNPV/Sf21 AcNPV/Sf9 AcNPV/High Five AcNPV/Sf9 BmNPV/Bm5 AcNPV/Sf9 AcNPV/Sf9 AcNPV/Sf21 AcNPV/Sf9 AcNPV/Sf21 AcNPV/Sf9 AcNPV/Sf9 AcNPV/Sf9

Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin Polyhedrin

>100 350 >500 50 250 200 >10 100–200 20 30–40 30 8–80 5–15

T-flask 4 l bioreactor Shake flask Spinner flasks 1.5 l bioreactor Spinner flasks Spinner flasks n.r. 15 l bioreactor n.r. Spinner flask 1 l bioreactor Spinner flasks

Luckow and Summers (1988) Caron et al. (1990) Wickham et al. (1992) Juarbe-Osorio (1993) Zhang et al. (1993) Lord et al. (1997) Coleman et al. (1997) Pittelkow et al. (1998) Cooley et al. (1998) Baldock et al. (2000) Meij et al. (2000) Pereira et al. (2001) Dorjsuren et al. (2003)

Secreted

Hu tPA Hu SEAP Rat PCE Hu IL-8 Hu LTA4H Hu C9 HIV gp120 Hu procolipase Hu PSA In JHE In JHE In JHE In JHE In JHE Hu IL-5 Ms OSF-2 Hu HCII GalNAc-transferase Hu IL-5 Hu LIF Hu ApoA-I Hu prorennin Ms FAK In transferrin Hu prorennin Ms GHBP Chk avidin Hu MMP9 Hu a1m Mn CatK Hu STC Hu GL Hu Apo A-I mutants Hu OPG-Fc Hu prolactin Hu prolactin In proPAP Hu MMP-9 Hu amylase In proPAP-2 Hu IL-3 Intracellular

CAT VP6 b-gal VP4 CAT Hu ACL NF6B1 and RelA Hu aminoacylase I Hu M/NEI Hu Syk EBNA1 Hu PARP KSHV Pol

Most reports are obtained from a survey of the Protein Expression and Purification journal (n.r., not reported; Hu, human; Ms, mouse; In, insect; Chk, chicken; Mn, monkey; tPA, tissue plasminogen activator; SEAP, placental secreted alkaline phosphatase; PCE, pancreatic cholesterol esterase; IL-8, interleukin-8; LTA4H, leukotriene A4 hydrolase; C9, complement protein 9; HIV gp120,

Insect Cell Culture and Recombinant Protein Expression Systems

485

Figure 2 One technique to improve the expression level of secreted proteins from baculovirus infected insect cells involves the removal of the signal peptide encoded by the gene and the creation of an artificial ATG at the start of the mature polypeptide coding region in the expression vector. This modification prevents the nascent polypeptide from entering the secretory pathway. In this example, the yield of biologically active juvenile hormone esterase (JHE) present in the culture supernatant of recombinant AcNPVinfected Sf21 cells is enhanced approximately 20-fold when the JHE signal peptide is removed (AcJHE-KKDsp) compared to the full length gene AcJHE-KK (Bonning et al., 1997). Left panel: Western blot analysis of 5 ml culture media to confirm the relative JHE expression levels in static cultures. Lane C, control culture medium; lane 1, day 6 cultutre medium form AcJHE-KKDsp infected Sf21 cells; lane 2, day 6 culture medium from AcJHE-KK infected Sf21 cells. Right panel: Batch production of JHE by AcJHE-KK or AcJHEKKDsp-infected Sf21 cells in serum-containing medium in 6-well plates over a 6-day period. Note that JHE produced by AcJHEKKDsp was not directed through the secretory pathway.

used membrane preparations from infected cells rather than the cells themselves to carry out their studies (Clawges et al., 1997). In addition, coupling of heterologous receptors with endogenous insect cell host proteins, such as G proteins, has generally produced low responses upon ligand binding (see below). This problem has been circumvented by coexpressing a GPCR with specific G protein subunits in order to assess agonist binding (Butkerait et al., 1995; Barr et al., 1997; Bouvier et al., 1998). 4.14.2.6.4. Compartmentalized proteins The BES has also been employed to synthesize and correctly process proteins targeted to subcellular compartments. A variety of mitochondrial proteins have been expressed using the BES, both membrane associated (Yet et al., 1995; Bader et al., 1998; Huang et al., 2000) and located within the mitochondrial matrix (Wang and Kaguni, 1999; Holcomb et al., 2000), although the latter reported incomplete

processing and inefficient targeting. Lysosomal enzymes have also been successfully expressed (Tschantz et al., 1999; Bromme and McGrath, 1996; Steed et al., 1998), as has a nucleolar protein (Ren et al., 1996) and a nuclear membrane protein (Bailer et al., 1995). 4.14.2.7. Other Applications of the Baculovirus Expression System

4.14.2.7.1. Virus-like particles The baculovirus expression system has proven to be very effective for the production of heterologous virus-like and core-like particles (VLPs and CLPs) destined for use in immunization or structural studies. Some examples include the poliovirus VLP (Urakawa et al., 1989), bluetongue virus CLP (French and Roy, 1990), Norwalk virus (Jiang et al., 1992), the papillomavirus (Cann et al., 1995), and herpes simplex virus (Newcomb et al., 1999). VLPs and CLPs can be architecturally complex and comprising several structural proteins in different molar

human immunodeficiency virus glycoprotein 120; PSA, prostrate specific antigen; JHE, Heliothis virescens juvenile hormone esterase; IL-5, interleukin-5; OSF-2, murine osteoblast specific factor 2; HCII, heparin cofactor II; GalNAc-transferase, bovine UDP-GalNAc polypeptide N-acetyl galactosaminyl transferase; LIF, leukemia inhibitory factor; Apo A1, apolipoprotein A-I; FAK, focal adhesion kinase; GHBP, growth hormone binding protein; MMP9 matrix metalloproteinase-9; a1m–a1, microglobulin; CatK, cathepsin K; STC, stanniocalcin; GL, gastric lipase; OPG-Fc, osteoprotegerin-immunoglobulin Fc fusion; proPAP, Manduca sexta prophenoloxidase-activating proteinase precursor; proPAP-2, M. sexta prophenoloxidase-activating proteinase-2 precursor; IL-3, interleukin-3; CAT, bacterial chloramphenicol acetyl-transferase; VP6, rotavirus VP6; b-gal, beta galactosidase; VP4, rotavirus outer capsid protein; ACL, ATP citrate lyase; NFkB1/RelA, subunits of the NF-kB transcriptional activator of immunoglobulin k light chain; M/NEI, monocyte/neutrophil elastase inhibitor; Syk, Syk protein; EBNAI, Epstein-Barr virus nuclear antigen 1; PARP, poly(ADP-ribose) polymerase; KSHV Pol, Kaposi’s sarcoma-associated herpesvirus DNA polymerase).

486 Insect Cell Culture and Recombinant Protein Expression Systems

gene encoding gp64 in the baculovirus genome, it was shown that foreign proteins (X) could be presented on the mature N-terminus of the duplicated X-gp64 (fusion) protein, in the envelope of BVs (Boublik et al., 1995). A demonstration of the feasibility of using baculovirus display for the screening of a library has been described (Ernst et al., 1998). Immunization of mice with purified virions displaying X-gp64 fusion proteins has also been useful for obtaining polyclonal and monoclonal antibodies against the antigen X (Lindley et al., 2000). This procedure eliminates one bottleneck in the conventional antibody production process of having to generate purified protein X for immunization. Due to an immunostimulatory effect often associated with viral immunogens (Minev et al., 1999), this method may be more effective than the conventional process for raising antibodies against certain antigens (Lindley et al., 2000).

Figure 3 A comparison of electron micrographs of native blue-tongue virus (BTV) virions with recombinant BTV viruslike particles (VLPs) produced and self-assembled in baculovirus-infected insect cells. For optimal synthesis of the BTV VLPs, a quadruple gene expression vector was used to coexpress BTV VP2, VP3, VP5, and VP7 proteins (Roy et al., 1997). At the top of the figure, a model of the BTV virion is shown. Images kindly supplied by Dr Polly Roy (London School of Hygiene and Tropical Medicine, UK).

proportions. It was observed that the coexpression of the various VLP structural proteins in baculovirusinfected insect cells results in the self-assembly of the VLP (Urakawa et al., 1989). Initially, the various structural proteins were produced by coinfection with individual recombinant baculoviruses. However, variation in the distribution of individual recombinant baculoviruses to each cell would result in differences in the quality of the VLP. Coexpression of proteins of the VLP or CLP using a single baculovirus with multiple promoters circumvents this problem (Figure 3; French and Roy, 1990; Belyaev et al., 1995). The ratio of each protein expressed can be controlled through the use of baculovirus promoters of different strength (Roy et al., 1997). 4.14.2.7.2. Surface display and antibody production The baculovirus virion has been developed as an eukaryotic alternative to the bacteriophage for the surface display of foreign proteins from an expression library and the selection of specific binding proteins (review: Grabherr et al., 2001). By duplicating the essential major envelope glycoprotein

4.14.2.7.3. Expression of recombinant proteins in insect larvae Insect larvae have occasionally been employed, instead of cultured insect cells, as hosts for recombinant baculoviruses for the expression of recombinant proteins (Maeda et al., 1985). Production in larvae can be inexpensive, high yielding, and easily scalable compared to cell culture. Yields around 1 mg per silkworm larva infected with recombinant BmNPV have been obtained for human interleukin-3 (Miyajima et al., 1987), hepatitis B surface antigen (Higashihashi et al., 1991), human CD66 antigens (Yamanaka et al., 1996), grass carp growth hormone (Ho et al., 1998), hepatitis E virus capsid protein (Sehgal et al., 2003), and human fibroblast growth factor (Wu et al., 2001). A bovine cardiac sodium– calcium exchanger has also been successfully expressed in T. ni larvae (Hale et al., 1999). However, the recovery of the expressed protein may be more problematic from larvae than from tissue culture because of increased proteolysis in the insect hemolymph and contamination of preparations by abundant hemolymph proteins. 4.14.2.7.4. Baculovirus transducing and transforming vectors Because of their ability to enter efficiently host and nonhost insect cells alike, baculovirus vectors can be designed with the goal of overexpressing regulatory factors and studying their role during development. Because baculoviruses are normally lethal to the host tissues, expression studies are by necessity limited to a brief period of experimentation extended between the initial infection and the activation of the host cell lysis process (Iatrou and Meidinger, 1989). For lepidopteran insect systems, this problem can be

Insect Cell Culture and Recombinant Protein Expression Systems

circumvented by the construction and use of mutant viruses that are unable to proceed through the late phases of the infection cycle. By contrast, infection of tissues of nonhost insects that are refractory to productive infection can result in more extended periods of expression and experimental observation (Oppenheimer et al., 1999). Two approaches have been utilized for the generation of transgenic lepidopteran insects using baculoviruses. One exploits the fact that AcNPV can infect nonhost lepidopteran insect larvae without killing the insect (Mori et al., 1995; Oppenheimer et al., 1999). Due to this property, the AcNPV can be used as a vehicle to efficiently deliver a transgene to germline cells, which is then targeted to chromosome sites for homologous recombination and generation of transgenic progeny (Yamao et al., 1999). A second approach uses a recombinant baculovirus with a deleted lef-8 gene (Iatrou et al., 2000). The lef-8 gene encodes a subunit of the viral RNA polymerase (Passarelli et al., 1994) that is necessary for mRNA transcription from the promoters of late and very late-phase baculovirus genes. Loss-of-function mutations in this gene prevent the progression from the replication phase to the virulent phase of the baculovirus infection cycle

f0020

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(Shikata et al., 1998). Accordingly, deletions of the lef-8 gene convert the baculovirus to a harmless, self-replicating extra-chromosomal entity (baculovirus artificial chromosome – BVAC), while host cells infected by it behave as normal ones (P.J. Farrell and K. Iatrou, unpublished results). Passage of the lef-8 deficient baculovirus through a rescuing cell line that constitutively expresses the Lef-8 protein enables the production of BVAC inoculum that maintains full infectivity for and replicating capacity in the host cells (Figure 4). The BVACs could potentially be used for prolonged protein expression by taking advantage of the high initial infection efficiency of cells by the baculovirus, the replication of the BVACs in them, and the subsequent mitotic transmission of the transgene-containing BVAC to the daughter cells. Furthermore, the prevention of both virus-induced damage to the secretory pathway and lysis would make BVACs more suitable for the expression of secreted and membrane proteins under the control of early viral or cellular promoters than conventional baculovirus expression vectors. Efforts are also being made to generate transgenic silkworms through the infection of germline cells with BVACs derived from BmNPV.

Figure 4 The potential for a nonlytic baculovirus is demonstrated. (a) Infection of normal insect cells by a wild-type baculovirus. The infected cells produce OBs (small, refractive objects) and eventually lyse. (b) A baculovirus artificial chromosome (BVAC) is created when the lef-8 gene is eliminated and replaced with a reporter gene expression b-galactosidase (dark cells). When the lef-8 gene product is supplied by a transformed cell line, the BVAC can complete the infection cycle that includes the formation of budded virions and occlusion bodies and cell lysis, in addition to the production of b-galactosidase. (c) The absence of the lef-8 gene product, in normal permissive cells infected by a BVAC, prevents completion of the baculovirus infection cycle. OBs are absent, the cells appear and replicate normally, and also express b-galactosidase.

488 Insect Cell Culture and Recombinant Protein Expression Systems

4.14.2.7.4. Transduction of mammalian cells Baculovirus virions were demonstrated to have the ability to enter certain cell lines derived from vertebrate species without evidence of viral gene expression (Volkman and Goldsmith, 1983; Carbonell and Miller, 1987). By incorporating mammalian expression cassettes into recombinant AcNPVs, several reports have appeared demonstrating that baculoviruses can serve as an efficient mode of gene transfer into a variety of primary and transformed mammalian cell lines, including those difficult to transfect using traditional methods, and express reporter proteins under the control of mammalian promoters (Hoffmann et al., 1995; Boyce and Bucher, 1996; Shoji et al., 1997; Yap et al., 1997; Condreay et al., 1999). Chromosomal integration and stable gene expression have also been demonstrated to be feasible by inclusion of a selectable marker (Condreay et al., 1999; Merrihew et al., 2001) or by integration signals from adenoassociated virus (AAV) into the baculovirus genome (Palombo et al., 1998). The advantages of using the baculovirus in this application include the high transfection efficiency (often greater than 90%; Condreay et al., 1999), low toxicity of the baculovirus to mammalian cells, and capacity of the baculovirus to carry large inserts. Recently, hybrids of the baculovirus and AAV were used to coinfect mammalian cells and produce high titer recombinant AAV for gene therapy applications (Sollerbrant et al., 2001).

conferring resistance to G418 (Jarvis et al., 1990), hygromycin B (Johansen et al., 1989), methotrexate (Shotkoski and Fallon, 1993), puromycin (McLachlin and Miller, 1997), and zeocin (Pfeifer et al., 1997) have all been used extensively with insect cell lines. The production of recombinant proteins in transfected or stably transformed insect cells provides considerable advantages over both the baculovirus expression system and transformed mammalian cells for the case of applications involving secreted and membrane-anchored proteins: insect cell lines are safe to humans, introns are spliced correctly and efficiently from expressed genomic DNAs and lysis does not occur, therefore allowing continuous, as opposed to batch-type, expression of the proteins and limiting proteolysis and facilitating purification of the secreted proteins. Furthermore, insect cells can perform most essential posttranslational modifications as efficiently as mammalian ones, and membrane proteins can be expressed in a stable physiological environment. Finally, most insect cell lines can grow in serum-free media to high densities, and many insect cell lines, particularly of lepidopteran origin, are already well characterized in large-scale suspension culture because of their role as hosts in the baculovirus expression system. Two major categories of plasmid-based expression systems have been developed utilizing promoters and cell lines derived from two different insect orders, Diptera and Lepidoptera. These are presented in more detail below.

4.14.3. Insect Cells as Hosts for Plasmid-Based Expression of Recombinant Proteins

4.14.3.2. Expression Systems Based on Dipteran Cell Lines

4.14.3.1. Introduction

Several baculovirus-free, plasmid-based approaches to recombinant protein expression in insect cells have been developed. In this technique, a plasmid expression cassette harboring a gene of interest is introduced into the host insect cell line using a variety of transfection techniques. The recombinant protein will be transiently expressed for a few days after transfection whereupon the cells or their media containing the recombinant protein can be harvested. Stable cell lines expressing the recombinant protein of interest, either continuously or by induction, can be generated by applying an antibiotic resistance-selection scheme over a period of several weeks. The antibiotic resistance gene can either be present on the same expression cassette as the gene of interest or supplied by cotransfection with a separate plasmid. Antibiotic selection schemes

Dipteran expression vectors mostly utilize the strong constitutive actin 5C promoter of D. melanogaster (Angelichio et al., 1991) or the inducible metallothionein promoter of Drosophila (Johansen et al., 1989; Kovach et al., 1992; Millar et al., 1995; Hegedus et al., 1998; Zhang et al., 2001) to drive foreign protein expression, in conjunction to a hygromycin B antibiotic selection scheme to generate a polyclonal population of Schneider 2 cell lines (S2 cells; Schneider, 1972), which takes approximately 3 weeks to accomplish (Johansen et al., 1989). The latter system has been developed by SmithKline-Beecham Pharmaceuticals and made commercially available through Invitrogen Corporation as the Drosophila Expression System (DESÕ). Drosophila S2 cells can grow in serum-free medium in batch suspension culture to cell densities up to 15  106 cells ml
1, although S2 cells are substantially smaller than most lepidopteran cell lines including Sf21, High FiveTM, and Bm5

Insect Cell Culture and Recombinant Protein Expression Systems

489

Table 2 Expression levels of some secreted proteins from stably transformed insect cells Protein

Level (mg l
1)

Culture conditions

Reference

S2 S2 S2 S2 S2 S2

22 2 5–35 1 3 2

n.r. n.r. n.r. n.r. n.r. Spinner flasks

Johansen et al. (1995) Culp et al. (1991) Ivey-Hoyle et al. (1991) Kirkpatrick et al. (1995) Denault et al. (2000) Shin and Cha (2003)

Sf9

1.0

Static

Jarvis et al. (1990)

Sf9 Sf9 Sf9 S2

0.5–1 10 18 10–15

Spinner flasks Spinner flasks Shake flasks Spinner flasks

Li et al. (2001) Hegedus et al. (1999) Pfeifer et al. (2001) Nilsen and Castellino (1999)

135–160 130–190 27–46

Static-spinner flask Static-spinner flask Spinner flask-static

Farrell et al. (1999) Farrell et al. (1998) Keith et al. (1999)

Cells

D. melanogaster metallothionein promoter

Hu IL-5 Modified HIV gp120 Modified HIV gp120 Hu IgG1 Hu SPC1 Hu EPO AcNPV ie-1 promoter

Hu tPA OpMNPV ie-2 promoter

Ms IgG1 Modified Hu p97 Modified Hu Factor X Hu plasminogen

B. mori cytoplasmic actin promoter, pIE1/153A vector

Hu tPA In JHE Hu GM-CSF

Bm5 Bm5 High Five

n.r., not reported; Hu, human; Ms, mouse; In, insect; IL-5, Interleukin-5; HIV gp120, human immunodeficiency virus glycoprotein 120; IgG1, immunoglobulin G1; SPC1, subtilisin-like proprotein convertase 1; EPO, erythropoietin; tPA, tissue plasminogen activator; p97, melanotransferrin; JHE, Heliothis virescens juvenile hormone esterase; GM-CSF, granulocyte macrophage colony stimulating factor.

cells. Enzymes, membrane receptors, ion channels, viral antigens, and monoclonal antibodies have been successfully produced using these systems (see Table 2). 4.14.3.3. Expression Systems Based on Lepidopteran Cell Lines

Lepidopteran expression vectors utilize a variety of promoters derived from baculoviruses and lepidopteran cells, including the AcNPV ie-1 promoter, the OpNPV ie-2 promoter, and the B. mori actin gene promoter; these promoters function mainly in lepidopteran cell lines such as Sf21, Sf9, High FiveTM, or Bm5 cells. 4.14.3.3.1. Constitutive expression 4.14.3.3.1.1. Baculovirus immediate early promoters A lepidopteran insect cell-based expression system was initially developed by Dr Don Jarvis at Texas A&M University (Jarvis et al., 1990). To generate stably transformed cell lines, Sf9 cells were cotransfected with a neomycin resistance plasmid, an expression vector employing the AcNPV immediate early gene promoter (ie-1) and a region containing mRNA polyadenylation signals (Guarino and Summers, 1987), followed by selection and isolation of G418 resistant clones over a period of 4 weeks following transfection. From the transformed insect cell clones, the expression levels of b-galactosidase (an intracellular protein) were approximately

100-fold lower than those obtained from a baculovirus expression vector employing the polyhedrin promoter, while the level of t-PA (a secreted protein) obtained was only two-fold lower. Despite the low values, these experiments demonstrated the potential of using stable lepidopteran cell lines for producing those recombinant proteins directed through the secretory pathway as an alternative to a baculovirus. The ie-1 gene promoter was also shown to function in dipteran cell lines (Vanden Broeck et al., 1995). Improvements in the expression vector were made by incorporating the AcNPV homologous repeat 5 (HR5) transcriptional enhancer element (Guarino et al., 1986) upstream of the ie-1 promoter (Jarvis et al., 1996). HR5 was shown to act in cis to significantly stimulate the expression of a reporter protein from early baculovirus promoters such as ie-1 and p35 in transient expression assays (Rodems and Friesen, 1993; Pullen and Friesen, 1995). A series of vectors utilizing the HR5-enhanced AcNPV ie-1 promoter are available from Novagen for stable or transient protein expression. A similar set of expression vectors has been developed based on the Orgyia pseudotsugata multicapsid nuclear polyhedrosis virus (OpMNPV) ie-2 promoter (Theilmann and Stewart, 1992; Pfeifer et al., 1997; Hegedus et al., 1998), which can function in both lepidopteran and dipteran insect cell lines. These vectors are available from Invitrogen as the Insect SelectTM vector set.

490 Insect Cell Culture and Recombinant Protein Expression Systems

4.14.3.3.1.2. Bombyx mori cytoplasmic actin promoter A powerful lepidopteran expression cassette, pIE1/153A, was recently developed in the laboratory of Dr Kostas Iatrou at the University of Calgary. The cassette utilizes the silk moth (B. mori) cytoplasmic actin gene promoter (Mounier and Prudhomme, 1986; Johnson et al., 1992), one of the stronger constitutive cellular promoters that is also active in a variety of transfected lepidopteran cell lines. Transcription from this cellular promoter was surprisingly found to be stimulated by the immediate early gene product (ie-1) of BmNPV, IE-1, a transcription factor capable of stimulating the in vitro rate of transcription from the actin promoter in trans by up to 100-fold (Lu et al., 1996). Stimulation of the actin promoter by up to two orders of magnitude was also obtained by linking in cis the homologous repeat 3 (HR3) region of BmNPV in various orientations relative to the actin promoter (Lu et al., 1997). Linkage of the ie-1 gene and the HR3 enhancer element with the actin gene promoter in the pIE1/153A expression cassette resulted in a stimulation of foreign gene expression directed by the actin promoter by approximately 5000-fold in transient expression assays for two reporter proteins (Lu et al., 1997). Stable cell lines have been generated by cotransfecting the cells with the expression cassette and a second plasmid conferring resistance to hygromycin B, puromycin, or G418. Reported expression levels of secreted proteins from stable cell lines transformed with this system have exceeded, by far, those obtained by the baculovirus expression system (Farrell et al., 1998; Farrell et al., 1999). Furthermore the expression plasmid functions in a wide variety of lepidopteran cell lines including those derived from B. mori, T. ni, Plodia interpuntella, L. dispar, Mamestra brassicae, C. fumiferana, and S. frugiperda (Keith et al., 1999). The expression cassette is also suited to scaled-up transient expression protocols that can yield tens of milligrams of recombinant protein per liter in 5 days posttransfection (Farrel and Iatrou, 2004) (Figure 5), thus avoiding the time consuming and labour intensive process of stable antibiotic selection and the cloning of high expressors. Derivatives of the pIE1/153A expression cassette are available from Dr Iatrou. 4.14.3.4. Insect Cell-Based Expression Using Inducible Promoter Systems

In contrast to constitutive promoters, the use of inducible promoters has the advantage that it allows the production of proteins that are toxic or growthinhibitory to the cells. In such cases, cells are first grown in batch mode (rather than continuously) to

Figure 5 Transient expression and affinity purification of histidine-tagged human secreted alkaline phosphatase (SEAP; kindly provided by Eric Carpentier and Amine Kamen, Biotechnology Research Institute, Montreal, Canada) from a lipofectinmediated transfection of High FiveTM cells in suspension culture using the pIE1/153A expression cassette. By comparison to a series of bovine serum albumin mass standards on the Coomassie blue-stained gel, approximately 50 mg ml
1 of SEAP was present in the culture supernatant 5 days after transfection and production in protein-free medium ESF-921 (lane marked ‘‘þ’’ compared to control marked ‘‘
’’). Two milligram of SEAP was recovered from one elution fraction (E) following Ni-ion affinity purification from 50 ml of culture supernatant.

high densities. When sufficiently high densities are reached, the inducer is added to the cell culture to induce high-level protein expression. Of importance to the choice of an inducible expression system is the induction ratio of gene expression. In the absence of inducer, promoter activity should be minimal as to prevent accumulation of toxic proteins during the growth phase of the cells. Addition of the inducer should, subsequently, result in the accumulation of high amounts of protein. 4.14.3.4.1. Heat shock promoter The Drosophila hsp70 promoter (Thummel and Pirrotta, 1992) has been widely used to achieve inducible gene expression in transgenic flies. The promoter is heat-inducible and functions during all stages of development and in all tissues (Steller and Pirrotta, 1984; Spradling, 1986). It has a low basal expression coupled with high inducibility (>100-fold at 42  C) (Huynh and Zieler, 1999). The promoter is not only functional in Drosophila cells but also in mosquito and lepidopteran cell lines (Zhao and Eggleston, 1999; Helgen and Fallon, 1990; Lan and Riddiford, 1997). While the hsp70 promoter has been used to mediate expression of membrane proteins (Shotkoski et al., 1996) and to drive expression of ‘‘helper’’ antibiotic resistance genes for the establishment of

Insect Cell Culture and Recombinant Protein Expression Systems

permanently transformed cell lines (Lycett and Crampton, 1993; Vanden Broeck et al., 1995) and transgenic mosquitoes (Miller et al., 1987), its application for the inducible expression of proteins at high levels for functional or structural characterization is considered limited. Long-term exposure to high temperature, which is considered necessary to reach the highest expression levels, is generally detrimental to the cell cultures. Also the translational induction to heat shock is reported to be much lower than the transcriptional induction (Cherbas et al., 1994), presumably because the heat shock interferes with posttranscriptional processing and translation. Heat shock (hsp70) promoter-based reporter and expression constructs were also incorporated in baculovirus vectors to monitor infection of tissues and cells and to achieve protein production. In the context of the baculovirus genome in infected cells, however, gene expression from the hsp70 promoter was constitutive rather than heat-inducible (Lee et al., 2000; Moto et al., 2003). 4.14.3.4.2. Metallothionein promoter Transcription of mammalian and Drosophila metallothionein genes is activated by heavy metal load via activation of the metal-responsive factor 1 (MTF-1) (Zhang et al., 2001). Inducible expression of genes via the metallothionein promoter is achieved in Drosophila S2 cells and other dipteran cell lines (Johansen et al., 1989; Kovach et al., 1992; Millar et al., 1995; Hegedus et al., 1998). Protein expression via the metallothionein promoter is induced by the addition of cadmium or copper ions (10–1000 mM) and induction levels of several 100-fold are achieved (Bunch et al., 1988; Otto et al., 1987; Zhang et al., 2001). The metallothionein promoter-based inducible system is considered suitable for the inducible expression of many different classes of proteins in dipteran cell lines. A disadvantage of the metal ion-inducible expression system is that at the high metal ion concentrations required for achieving the highest expression levels, toxic effects are also observed on the cells, such as cytoplasmic vacuolization and granule formation as well as an associated reduction in their growth rate (Bunch et al., 1988). Thus, optimal protein production requires determination of the right balance between degree of induction of gene expression and general cellular toxicity. While the metal ion-mediated induction works efficiently in Drosophila and mammalian cell lines (Zhang et al., 2001), in lepidopteran cell lines such as Sf9, Ld652Y, and Bm5, the metallothionein

491

promoter was either not or only marginally functional (Hegedus et al., 1998; V. Douris, L. Swevers and K. Iatrou, unpublished observations). 4.14.3.4.3. Ecdysone-inducible promoters Many insect tissue culture cell lines are sensitive to the insect molting hormone 20-hydroxyecdysone (20E) and respond to it by morphological changes and a decline in growth rates (Sorbrier et al., 1989; Sohi et al., 1995; Smagghe et al., 2003). At the molecular level, 20E exerts its actions by binding to the ecdysone nuclear receptor (EcR) that exists as a heterodimer with the nuclear receptor Ultraspiracle (USP) (Thomas et al., 1993; Yao et al., 1993; Swevers et al., 1996) and the hormone-bound complex directly activates transcription at ecdysone-response elements (EREs) in promoters of target genes (Antoniewski et al., 1996; Vogtli et al., 1998; Wang et al., 1998; see Chapter 3.5). Early-response genes, which are directly regulated by the EcR/USP complex, are induced in insect cell lines after challenge with 20E (Sohi et al., 1995; Chen et al., 2002; Swevers et al., 2003b), indicating that all transduction elements for gene activation by 20E are present. Basal reporter cassettes that contain multiple repeats of an ERE derived from the Drosophila hsp27 promoter are 1000- to 2000-fold induced by mM quantities of 20E in Drosophila S2 cells as well as Bombyx-derived (lepidopteran) Bm5 cells (Koelle et al., 1991; Swevers et al., 2003b). The induced expression levels are comparable to those obtained by strong constitutive promoters (Swevers et al., 2003b) and silk moth (lepidopteran) transformed cell lines incorporating an ecdysteroid-inducible expression system have been described for production of recombinant proteins (Tomita et al., 2001) as well as high-throughput screening for potential 20E agonists and antagonists (Swevers et al., 2003b). An advantage of the ecdysteroid-inducible expression system is that, in the absence of ligand, the EcR/USP heterodimer functions as a repressor of gene transcription (Tsai et al., 1999) and the basal expression levels in the absence of the inducer are very low. Because 20E is a natural hormone, toxicity effects on the cell cultures are virtually nonexistent. Nevertheless, it remains to be investigated in detail whether the physiological effects of 20E on the function of the cells could potentially interfere with their capacity for protein production. It is expected that this will not be the case: using the baculovirus/lepidopteran cell line expression system, it was reported that the addition of ecdysteroids actually increased recombinant protein production (Sarvari et al., 1990).

492 Insect Cell Culture and Recombinant Protein Expression Systems

4.14.3.5. Classes of Recombinant Proteins Expressed by Transformed Insect Cells

4.14.3.5.1. Secreted proteins A variety of heterologous secreted proteins have been expressed from stable insect cell lines at similar or higher expression levels than can be obtained using baculovirus expression vectors (Table 2). Certainly the cellular or early-phase baculovirus promoters employed are not as transcriptionally powerful as late- and very late phase promoters such as those of the polyhedrin, p10 or basic protein genes. However, the fact that the cell’s secretory pathway is not being damaged in the absence of viral infection, contributes to the improvement in the expression level and quality of the overexpressed protein compared to expression using recombinant baculovirus. The substitution of the native mammalian signal peptide coding sequence in a heterologous mammalian gene with a sequence encoding an insectspecific signal peptide was found to have no effect on the resulting expression level in transfected insect cells (Farrell et al., 2000). 4.14.3.5.2. Membrane proteins As with secreted proteins, the absence of viral infection may provide a superior cellular environment for the production of membrane proteins which also traverse the secretory pathway. Several functional membrane proteins have been successfully expressed in stable insect cell lines including ion exchangers (Szerenscei et al., 2000), transmitter gated ion channels (Joyce et al., 1993; Millar et al., 1994; Smith et al., 1995; Atkinson et al., 1996; Buckingham et al., 1996), and GPCRs (Kleymann et al., 1993; Swevers et al., 2003a; see Chapter 5.5). As found with the baculovirus expression system (Bouvier et al., 1998), coupling of heterologous GPCRs with endogenous insect G proteins may be inefficient following ligand binding, even in transformed insect cells (Figure 6). However, a strategy that has proven useful for the functional expression of heterologous GPCRs is the coexpression of mammalian G proteins in stably transformed insect cells (L. Swevers, K. Iatrou, E. Morou, N. Balatsos, and Z. Georgoussi, unpublished data; see also further below). 4.14.3.5.3. Intracellular proteins The use of transformed insect cells for the expression of intracellular proteins is of limited potential compared to the baculovirus expression system. There are a few reports on the concentrations of intracellular proteins obtained, however the promoters employed in stable gene expression from insect cells are considerably weaker than those utilized in baculovirus

Figure 6 Fluorescent spectrophotometer outputs of calcium signaling assays to compare the G protein-coupled receptor (GPCR) response of a mammalian GPCR, the rat protease activated receptor 2 (rPAR2; Hollenberg et al., 1996), expressed in stably transformed insect (panel a, High FiveTM) and mammalian (panel b, KNRK) cell lines. The amplitude reflects elevations in intracellular calcium with time, following exposure to the rPAR-2 peptide agonist, SLIGRL, at the time points denoted by the filled square and circle. Despite the fact that the receptor density was higher in the insect cell line (data not shown), the results imply that this mammalian GPCR does not couple well with the endogenous insect G proteins. Courtesy of Dr Morley Hollenberg, Mamoud Saifeddine and Bajhat Al-Ani (University of Calgary, Canada).

expression vectors (Jarvis et al., 1990; Percival et al., 1997; Pfeifer et al., 1997). Furthermore, the purification of heterologous intracellular proteins, expressed at relatively low levels inside insect cells, from endogenous cellular protein mixtures is expected to be difficult. Secretion of the intracellular protein would facilitate the purification process. However, the mere fusion of an insect-specific signal peptide to the N-terminus of an intracellular protein failed to provide the means necessary to secrete intracellular proteins (Farrell et al., 2000). It was realized that other biological signals in addition to a signal peptide are required for efficient secretion from insect and mammalian cells into an extracellular environment. These could be supplied by fusing the complete coding sequence of a secreted protein to that of an intracellular protein (Figure 7; Farrell et al., 2000). The chimeric proteins produced are soluble and can be easily purified from

Insect Cell Culture and Recombinant Protein Expression Systems

Figure 7 Secretion of luciferase from lepidopteran insect cells transfected with the pIE1/153A expression plasmid that encompasses a luciferase secretion module. Normally, luciferase (LUX) is expressed intracellularly, however, by fusing it with a secreted protein, in this case the human granulocyte macrophage colony stimulating factor (hGMCSF), the normally intracellular protein can be dragged into the supernatant as a chimera. High Five cell culture supernatants are shown in the SDS-PAGE/Western blots probed with (left) an anti-hGMCSF antibody and (right) an anti-LUX antibody. The lane marked ‘‘C’’ contains protein from a control transfection using the pIE1/153A expression vector without inserted transgene. Lanes 1 contain supernatants from cells transfected with vector pIE1/153.lux directing intracellular expression of luciferase, and Lanes 2 contain supernatants from cells transfected with vector pIE1/153A.hgmcsf-lux directing the secretion of the overexpressd hGMCSF-LUX fusion protein. Although some luciferase does leak from the cells naturally, a large proportion is secreted when expressed as a hGMCSF-LUX fusion protein.

the cell culture supernatant under nondenaturing conditions.

4.14.4. Insect Cell-Based High-Throughput Screening Systems In cell-based detection systems, cell lines are engineered such that they respond to a biological stimulus by the generation of an easily detectable, fluorescent or luminescent, signal. Accordingly, cell-based detection systems integrate two relevant genetic elements, (1) an expression element that directs expression of the target molecule (e.g., a receptor) thus allowing the cells to respond specifically to bioactive compounds (e.g., ligands) that interact specifically with the target and initiate a downstream cell signaling response, and (2) a reporter element that allows the cells to generate an easily measurable response (fluorescent or luminescent signal) following activation of the expressed receptor by the bioactive compound. Insect cell-based high throughput screening systems have been developed for two major types of receptors, nuclear hormone receptors (see Chapters 3.4–3.6) and membrane-anchored GPCRs (see Chapter 5.5). Because nuclear hormone

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receptors activate directly transcription at specific target sites in the DNA (Gronemeyer and Laudet, 1995), detection of nuclear receptor activation occurs by induction of expression of appropriate fluorescent (GFP or its variants) or luminescent (luciferase) reporter genes. On the other hand, GPCRs, which are subdivided into several classes according to the type of signal transduction pathway that they generate at the plasma membrane (cAMP up- or downregulation, release of Ca2þ, activation/inhibition of ion channels) (Hamm, 2001; Marinissen and Gudkind, 2001) require different reporter assays that are designed according to the specificity of each receptor class. Since the release of Ca2þ can be easily measured by fluorescent (e.g., employing the Ca2þ-sensitive fluorescent dyes fura-2 or fluo-3) or bioluminescent (coelenterazine/aequorin-based) methods (Grynkiewicz et al., 1985; Knight et al., 1991), activation of GPCRs that naturally function by Ca2þ release can be monitored by the detection of Ca2þ accumulation. For other GPCRs, the possibility exists to coexpress them with the mammalian Ga16 protein, which can couple GPCRs of other transduction classes to the Ca2þ pathway (Kostenis, 2001). 4.14.4.1. Nuclear Receptors: The Ecdysone Receptor

Of the approximately 20 nuclear hormone receptors identified in the insect (Drosophila or Anopheles) genome (Adams et al., 2000; Zdobnov et al., 2002; see Chapters 3.5 and 3.6), only one has a clearly identified ligand. This is EcR, the receptor for the insect moulting hormone 20E (Escriva et al., 1997). Because many insect cell lines express functional EcR receptors (Sohi et al., 1995; Chen et al., 2002; Swevers et al., 2003a), they have the potential to be developed as screening systems for the 20E hormone (see Chapter 3.4). A microplate-based bioassay, for the detection of 20E mimetic (agonistic and antagonistic) activities, was described that is based on changes in growth or morphology of the Drosophila BII tumorous blood cell line (Clement et al., 1993). This cell line is used to screen natural products from plants and fungi as well as collections of synthetic ecdysteroid derivatives for moulting and antimoulting hormone activity (Dinan et al., 1997; Harmatha and Dinan, 1997; Dinan et al., 2001; Harmatha et al., 2002). Another, more recently developed high throughput screening system for 20E agonists and antagonists, is based on silk moth-derived Bm5 cells that have been engineered to incorporate an ecdysteroidresponsive GFP reporter cassette in their genome

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thus validating the usefulness of the fluorescent screening system for rapid selection of 20E mimics as potential new insecticides. 4.14.4.2. G Protein-Coupled Receptors

Figure 8 Establishment of a high-throughput screening system for ecdysone agonists and antagonists based on transformed silk moth-derived Bm5 cells. (a) Schematic presentation of the ecdysone-responsive reporter constructs. EcRE, ecdysone-responsive element; CAT, chloramphenicol acetyl transferase; GFP, green fluorescent protein. (b) Doseresponse curve of the ecdysone-responsive CAT reporter construct in transient expression experiments. Shown is the fold inducibility in function at different hormone concentrations. (c) Induction of green fluorescence by 20E in a clonal transformed cell line that contains integrated copies of the ecdysone-responsive GFP reporter construct in their chromosomes.

(Swevers et al., 2003b; Figure 8). The half-maximal response of activation of the reporter cassette is elicited by concentrations of 20E between 50 and 100 nM, which are comparable to the concentrations required for induction of morphological or growth inhibitory responses in other cell lines (Cherbas et al., 1980; Dinan et al., 1997). An important advantage of the ecdysteroid-responsive fluorescent cell lines is that the amount of fluorescence emitted by the cells can be easily detected and quantified from cells seeded in multiwell plates by a fluorescence plate reader, thus rendering the system amenable to high-throughput analysis. The ecdysteroid-inducible fluorescent Bm5 cell lines were used successfully to screen a collection of plant extracts as well as a chemical library of potential nonsteroidal agonists (dibenzoyl hydrazine derivatives) for detection of 20E agonist and antagonist activities (Swevers et al., 2003b). The dibenzoyl hydrazine compounds selected by the high-throughput assay were also effective in larval toxicity tests,

The majority of signal transduction by hormones or neurotransmitters is mediated by GPCRs (Vanden Broeck, 2001). In the two insects whose complete genome sequence is known, D. melanogaster and Anopheles gambiae, GPCRs comprise the largest number of genes, accounting for 1–2% of the total gene number (Hill et al., 2002). Similarly, in the human genome more than 600 GPCR genes have been identified. Human GPCRs are considered important targets for drug development, and it is estimated that up to 40–50% of the available drugs are modulators of GPCR function (Kostenis, 2001). Using the AcNPV-based baculovirus expression system, several types of mammalian GPCRs have been successfully expressed in S. frugiperda Sf9 and T. ni Hi5 cell lines (Vasudevan et al., 1992; Ng et al., 1995, 1997; Zhang et al., 1995; Wehmeyer and Schulz, 1997; Ohtaki et al., 1998). While the AcNPV-based expression system is not amenable to high throughput screening development because the cells are lysed at the end of the infection cycle, these studies nevertheless demonstrated that the receptors that were overexpressed were functional and could couple efficiently to downstream signaling elements such as ion channels and the adenylate cyclase system. Thus, using plasmidbased expression systems that do not result in cell lysis (Farrell et al., 1998), transformed cell lines can be obtained that express constitutively functional GPCRs for screening of ligand mimetics. Drosophila Schneider 2 cells have been used by several research groups to achieve permanent and functional expression of insect as well as mammalian GPCRs (Joyce et al., 1993; Kleymann et al., 1993; Millar et al., 1995; Tota et al., 1995; Vanden Broeck et al., 1995; Buckingham et al., 1996; Torfs et al., 2000; Perret et al., 2003). In the case of a tachykinin receptor from the stable fly Stomoxys calcitrans, activation resulted in the detection of Ca2þ release by fluorescent and luminescent methods, thus making the transformed cell line an effective system for the screening of tachykinin-like ligands (Torfs et al., 2002). In mammalian cell-based screening systems, the ‘‘promiscuous’’ Ga proteins, Ga15 and Ga16, have been used to couple the activation of any GPCR to the Ca2þ signaling pathway (Stables et al., 1997; Kostenis, 2001). The presence of the Ga16 protein results in the activation of phospholipase Cb

Insect Cell Culture and Recombinant Protein Expression Systems

(PLCb) resulting in the production of the secondary messengers diacyl glycerol (DAG) and inositol (1,4,5)-triphosphate (IP3). While DAG activates other downstream pathways through activation of protein kinase C, the action of IP3 results in the release of Ca2þ from intracellular stores (Stables et al., 1997), which can be subsequently detected easily by fluorescent or bioluminescent methods (Grynkiewicz et al., 1985; Knight et al., 1991). Cell lines overexpressing Ga16 protein can, therefore, be used to screen for ligands of GPCRs that normally couple to other second messenger or to ion channels. This strategy has been employed recently in the case of transformed silk moth Bm5 cells that overexpress the murine d opioid receptor: coexpression of the human Ga16 protein in these cells has resulted in the stimulation of the coupling of the mouse receptor to the PLCb/IP3 pathway, thus making the transformed cell line a very effective screening system for ligands of the d opioid receptor (L. Swevers, K. Iatrou, E. Morou, N. Balatsos and Z. Georgoussi, unpublished data).

4.14.5. Glycosylation of Recombinant Proteins in Insect Cells Considerable attention has been paid to the glycosylation patterns of heterologous recombinant proteins produced in insect cells, particularly N-linked glycosylation. Several excellent and comprehensive reviews in this area have been published by Jarvis et al. (1998), Altmann et al. (1999), Marchal et al. (2001), and Jarvis (2003). In these reviews, it is agreed that oligomannose (five to nine mannose residues; Altmann et al., 1999) and fucosylated trimannose type carbohydrate (Jarvis et al., 1998) moieties dominate the N-linked glycosylation pattern of heterologous secreted and membrane proteins expressed in insect cells. The lack of complex glycosylation, such as the presence of terminal sialic acid found on many mammalian glycoproteins, excludes the use of insect cells to express mammalian glycoproteins for some applications. One solution to this limitation is to engineer baculoviruses (Jarvis et al., 1996; Wagner et al., 1996) or insect cell lines (Hollister et al., 1998) to express mammalian glycotransferases necessary for complex glycosylation of an over-expressed protein. The coexpression of two mammalian glycotransferases, b-1,4-galactosyltransferase and a-2,6-sialyltransferase, enabled an insect cell for the first time to produce an artificially sialylated baculovirus glycoprotein gp64 (Seo et al., 2001). This research will almost certainly extend the utility of insect

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cells to the production of ‘‘mammalized’’ complex glycoproteins.

4.14.6. Conclusions and Future Perspectives Baculoviruses and insect cell lines have acquired a significant biotechnological niche for the production of proteins of general scientific and pharmaceutical value. While baculovirus vectors provide a superior system for high level expression of intracellular proteins, their use for the production of proteins that require an intact cellular environment is considered to be suboptimal. In contrast, the recently developed expression systems that employ baculovirus-derived genetic elements and direct continuous high level expression in transformed insect cells, not only circumvent the major drawbacks of the baculovirus expression systems for secreted and membrane-anchored proteins but can also be employed as high-throughput screening tools for the identification of bioactive substances with defined biological specificities. For the baculovirus system, great technical improvements have occurred, mostly with regard to the efficiency and ease of generating recombinant viruses, with the best case being the generation of recombinant viruses through direct cloning into bacterially isolated bacmid chromosomal DNA. Because of the high degree of sophistication already achieved, relatively little improvement in this system is expected in the coming years. By contrast, the use of plasmid-based expression systems for insect cells is relatively recent and important improvements and new applications are expected to arise in the future. Plasmid-based expression systems are not, in general, considered optimal for the production of intracellular proteins, because their isolation requires cell lysis and purification from a multitude of contaminating cellular proteins. Improvements in the production process are expected to include the redirection of normally intracellular recombinant proteins toward the extracellular medium, from which they can be purified with relative ease, particularly if the medium is serum-free. To test this strategy, secretion modules derived from secreted proteins, such as JHE and granulocytemacrophage colony-stimulating factor (GM-CSF), have been fused in-frame with the N-termini of intracellular proteins, such as the CAT enzyme and the BmCF1 nuclear receptor (Farrell et al., 2000). Efficient secretion to the extracellular medium has been reported and, because the engineered proteins also contained a histidine tag and an enteropeptidase cleavage site, single-step purification

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could be achieved by metal-ion affinity binding coupled with proteolytic cleavage (Farrell et al., 2000). Future modifications in this method may include the use of alternative affinity tags and proteolytic sites as well as improved versions of the secretion modules. More recently, Drosophila S2 cell lines have been used as expression hosts for the purification of multiprotein complexes that are dedicated to the execution of particular cellular functions (Forler et al., 2003). Key factors involved in particular cellular processes (e.g., splicing of mRNA precursor molecules, RNA quality control, nuclear export) were expressed as double-tagged proteins, while the expression of endogenous untagged proteins was suppressed by RNA interference. Tagged proteins, together with assembled interacting factors, were purified by the ‘‘tandem-affinity purification’’ strategy and component proteins of purified complexes were identified by Western blot and mass spectrometry (Forler et al., 2003). The technique will be valuable for the characterization of multiprotein functional complexes operating in metazoan species, in general, and in insect ones, in particular. Transformation of insect cell lines with expression constructs still represents a cumbersome process requiring selection of resistant cell clones after addition of antibiotics. It is expected, however, that alternatives to the method of selecting cells containing genomic integrants of plasmid vectors by antibiotic selection, including the use of vectors capable of directing autonomous integration into the host genome, will be developed in the immediate future. In fact, recent reports suggest that the use of a plasmid encompassing the genomic sequences of the Junonia coenia densovirus (JcDNV) directs autonomous genomic integration in insect cells grown in culture, as well as in vivo, without affecting the normal physiology of the host cells in any detectable way (Bossin et al., 2003). Furthermore, it has also been reported that this system offers the additional advantage of integrated copy number manipulation through directed deletion of viral genome sequences from the expression plasmid (Bossin et al., 2003). Although the production capabilities of this system have yet to be tested with specific protein models, it is anticipated that the incorporation in it of promoter systems that are in current use as drivers of recombinant protein expression in insect cells, will expand further the current capabilities and advantages of the nonlytic systems utilizing insect cells as hosts for continuous high level production of recombinant proteins over lytic ones. The major advantage of the use of the densovirus-based transformation system is the ease of selection of transformed cells

by FACS analysis through the detection of red fluorescence in the cell nuclei (achieved by the nuclear expression of DsRed/viral capsid protein fusions directed by the use of the viral P9 promoter; Bossin et al., 2003). Finally, the detection of efficient densovirus amplification and densovirus-derived transgene expression in Drosophila larval and adult tissues (Royer et al., 2001) indicates that the densovirus-based transformation system is not limited to lepidopteran cell lines but can be applied to dipteran cell lines as well. In mammalian systems, many nuclear receptors and GPCRs have been characterized at the molecular level using transformed cell lines, and this has led to the discovery of lead compounds (and drugs) that interfere specifically at very low doses with the function of the receptors. Equivalent data on insect receptors, however, are currently lacking to a great extent. Although the identification of the natural ligands for orphan insect (Drosophila) GPCRs is now occurring at a good pace, the identification of the relevant ligands was made possible through the use of heterologous expression systems such as transformed mammalian cell lines or Xenopus oocytes (Birgu¨ l et al., 1999; Meeusen et al., 2002; Staubli et al., 2002; Johnson et al., 2003). In this regard, we expect that insect cell lines would provide better host expression systems for insect GPCRs and could be employed for a more effective screening for compounds that interfere with their functions. The same applies for the identification of ligands for the approximately 20 orphan nuclear receptors that exist in the insect genome (Adams et al., 2000; Zdobnov et al., 2002). The transcriptional properties of several of these receptors have been characterized (e.g., the HNF-4 receptor (Kapitskaya et al., 1998), the FTZ-F1 receptor (Suzuki et al., 2001), and the HR3 and E75 receptors (Swevers et al., 2002)) and it should be relatively straightforward to design cell-based reporter systems for screening of ligands, particularly small molecules capable of blocking their functions. In this regard, it is worth noting that the Drosophila DHR38/USP receptor complex was recently found to be activated by various ecdysteroids (Baker et al., 2003), illustrating the as yet unrealized potential of orphan nuclear receptors as targets for interference by small molecule ligands. Thus, like their mammalian counterparts, insect orphan receptors can be considered as targets for interference by small molecules, which can be developed into environmentally friendly insecticides that function as ‘‘endocrine disruptors,’’ and insect cell-based expression systems are anticipated to play an instrumental role in the discovery effort.

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5.1 Sodium Channels D M Soderlund, Cornell University, Geneva, NY, USA ß 2005, Elsevier BV. All Rights Reserved.

5.1.1. Introduction 5.1.1.1. Function and Structure of Voltage-Sensitive Sodium Channels 5.1.1.2. Sodium Channels as Targets for Neurotoxicants 5.1.1.3. Progress in Insect Sodium Channel Research Since 1985 5.1.2. Sodium Channel Genes in Insects 5.1.2.1. Sodium Channel Genes in Drosophila melanogaster 5.1.2.2. Sodium Channel Genes in Other Insect Species 5.1.3. Functional Expression of Cloned Insect Sodium Channels 5.1.3.1. Expression of Functional Sodium Channels in Xenopus Oocytes 5.1.3.2. Pharmacological Properties of Expressed Insect Sodium Channels 5.1.3.3. Functional Characterization of Sodium Channel Splice Variants 5.1.4. Sodium Channels and Knockdown Resistance to Pyrethroids 5.1.4.1. Knockdown Resistance 5.1.4.2. Altered Sodium Channel Regulation as a Mechanism of Knockdown Resistance 5.1.4.3. Genetic Linkage between Knockdown Resistance and Sodium Channel Genes 5.1.4.4. Identification of Resistance-Associated Mutations 5.1.4.5. Functional Analysis of Resistance-Associated Mutations 5.1.5. Conclusions 5.1.5.1. Unique Features of Insect Sodium Channels 5.1.5.2. Sodium Channels and Pyrethroid Resistance 5.1.5.3. Future Exploitation of the Sodium Channel as an Insecticide Target

5.1.1. Introduction 5.1.1.1. Function and Structure of Voltage-Sensitive Sodium Channels

Voltage-sensitive sodium channels mediate the transient increase in sodium ion permeability that underlies the rising phase of the electrical action potential in most types of excitable cells (Hille, 2001). In vertebrates, sodium channels are found in neurons and cardiac and skeletal muscle cells as well as glial and neuroendocrine cells (Goldin, 2002). In contrast, voltage-sensitive sodium channels in insects appear to be limited in distribution to neurons (Littleton and Ganetzky, 2000). The principal structural element of voltage-sensitive sodium channels is a large (
260 kDa) a subunit that forms the ion pore and confers the functional and pharmacological properties of the channel (Catterall, 2000; Yu and Catterall, 2003). Sodium channel a subunits are pseudotetrameric proteins that contain four internally homologous domains, each of which contains six hydrophobic transmembrane helices and additional hydrophobic segments that contribute to the formation of the ion pore (Figure 1). The four internally homologous domains form a radially symmetrical assembly with

1 1 2 4 4 4 7 9 9 9 10 11 11 11 11 12 14 17 17 17 18

the ion pore at the center (Guy and Seetharamulu, 1986; Yellen, 1998; Lipkind and Fozzard, 2000). Voltage-sensitive sodium channel a subunits are part of a larger family of cation channels that also includes cyclic nucleotide-gated channels and voltage-sensitive potassium and calcium channels (Strong et al., 1993). Whereas voltage-sensitive sodium and calcium channels share the pseudotetrameric four-domain structure shown in Figure 1, voltage-sensitive potassium channels and cyclic nucleotide-gated channels are composed of four separate subunits, each of which corresponds to a single domain of sodium and calcium channels. The phylogenetic distribution of voltage-sensitive cation channels and amino acid sequence comparisons among and between channel classes suggest that the earliest channels were tetramers of four separate subunits, like voltage-sensitive potassium channels (Strong et al., 1993; Goldin, 2002). These data also suggest that four-domain calcium channels arose from such channels via two rounds of intragenic duplication. The four-domain voltage-sensitive sodium channels, which appear later in evolution than calcium channels, are likely to have arisen from duplication and divergence of voltage-sensitive calcium channels.

2 Sodium Channels

Figure 1 Diagrammatic representations of the structure of vertebrate and insect sodium channel subunits. (a) Sodium channel a subunit, showing the four internally homologous domains (I–IV), each containing six hydrophobic transmembrane helices. Also illustrated are other structural elements relevant to sodium channel function: the multiple positively charged residues (þ) in the four S4 helices that constitute the voltage sensor, the four amino acids constituting the selectivity filter (the ‘‘DEKA motif’’) located in the pore-forming regions of the four internally homologous domains, and the inactivation gate peptide between the third and fourth domains. (b) Vertebrate sodium channel auxiliary b subunit. (c) Insect sodium channel auxiliary tipE subunit.

Further sodium channel gene duplication and divergence in the course of mammalian evolution has resulted in at least 10 sodium channel a subunit genes that differ in amino acid sequence, developmental and anatomical distribution, and biophysical and pharmacological properties (Goldin et al., 2000; Goldin, 2002). In mammalian brain and skeletal muscle, sodium channel a subunits are associated with one or more smaller auxiliary b subunits that modulate the expression and functional properties of sodium channel a subunits (Catterall, 2000; Goldin, 2001).

5.1.1.2. Sodium Channels as Targets for Neurotoxicants

Voltage-sensitive sodium channels are the site of action of a wide structural variety of naturally occurring neurotoxins that contribute to the chemical ecology of predation and defense (Ceste`le and Catterall, 2000; Blumenthal and Seibert, 2003; Wang and Wang, 2003). These sites, together with binding sites for synthetic neurotoxicants and drugs, identify at least 10 distinct binding domains associated with the voltage-sensitive sodium channel.

Sodium Channels 3

Five principal neurotoxin recognition sites, designated as sites 1–5 (Catterall, 1988, 1992), have been identified in both functional assays and in radioligand binding experiments (see Table 1). These sites are thought to represent physically nonoverlapping domains of the sodium channel protein that interact allosterically as the channel protein changes conformation in response to the binding of one or more neurotoxins. Site 1 binds the watersoluble toxins tetrodotoxin (TTX) and saxitoxin (STX), which interact at or near the extracellular opening of the ion pore and block ion transport through the channel. Site 2 binds a structurally heterogeneous group of lipophilic toxins that alter both the opening (activation) and closing (inactivation) of sodium channels. Site 2 neurotoxins shift the voltage dependency of sodium channel activation toward more negative membrane potentials, thereby increasing the probability that channels will open at normal membrane resting potentials. These toxins also slow or completely block sodium channel

Table 1 Identified and inferred binding domains on the voltage-sensitive sodium channel Sitea

Active neurotoxinsb

Physiological effect

Binding domains identified with specific radioligands

1

2

3 4 5 6 7

Tetrodotoxin Saxitoxin m-Conotoxin Veratridine Batrachotoxin Aconitine Grayanotoxins Pumiliotoxin-B N-alkylamides a Scorpion toxins Sea anemone toxins b Scorpion toxins Brevetoxins Ciguatoxin d-Conotoxins Conus striatus toxin DDT and pyrethroids

Inhibit ion transport

Persistent activation

Prolong inactivation Enhance activation Persistent activation Prolong inactivation Prolong inactivation

Binding domains inferred but not characterized with radioligands

8 9 10

mO-conotoxins Gonioporatoxin Local anesthetics Anticonvulsants Antiarrhythmics Pyrazolines

Inhibit ion transport Prolong inactivation Inhibit ion transport

a Sites 1–5 after Catterall (1988); sites 6 and 7 after Zlotkin (1999); sites 8–10 are assigned arbitrarily to distinguish them from sites 1–7. b Insecticides are capitalized.

inactivation, thereby prolonging the open state of the channel. Site 3 binds a group of polypeptide toxins, called a-toxins, isolated from scorpion venoms or sea anemone nematocysts. These toxins selectively prolong sodium channel inactivation without affecting the rate or voltage dependency of channel opening and allosterically enhance the action of compounds acting at site 2. Site 4 binds a second group of polypeptide scorpion toxins, called b-toxins, which selectively enhance sodium channel activation and do not interact allosterically with site 2 compounds. Site 5 binds the brevetoxins and ciguatoxin, lipophilic compounds isolated from marine dinoflagellates, which shift the voltage dependency of sodium channel activation and prolong inactivation in a manner similar to site 2 neurotoxins. However, these compounds bind at a site distinct from site 2 and interact allosterically with compounds acting at site 2. Subsequent research has identified two additional binding domains that are labeled with specific radioligands and shown to be distinct from sites 1 to 5. Site 6 (Table 1) binds d-conotoxins and Conus striatus toxin (Gonoi et al., 1987; Fainzilber et al., 1994). Site 7 (Table 1) (also designated site 6 in some classifications) binds dichlorodiphenyltrichloroethane (DDT) and pyrethroids (Bloomquist and Soderlund, 1988; Lombet et al., 1988; Trainer et al., 1997). At least three other binding domains (arbitrarily designated as sites 8–10 in Table 1) have been postulated to account for the actions on sodium channels of mO-conotoxins (Terlau et al., 1996), Goniopora coral toxin (Gonoi et al., 1986), and local anesthetics, class I antiarrhythmics, and class I anticonvulsants (Catterall, 1987). Sodium channels are widely recognized as the principal target site for diphenylethane (e.g., DDT) and pyrethroid insecticides (Sattelle and Yamamoto, 1988; Bloomquist, 1993; Soderlund, 1995; Narahashi, 1996). In addition, other toxicantbinding domains on the sodium channel are important target sites for the continued discovery and development of novel insect control agents. Two additional classes of synthetic insecticides alter sodium channel function by binding to sites that are distinct from the pyrethroid recognition site. Synthetic N-alkylamide insecticides, which are analogs of insecticidal natural products, produce excitatory effects by binding to site 2 (Ottea et al., 1989, 1990), whereas pyrazolines (also called dihydropyrazoles) and structurally related insecticides suppress normal nerve activity by the voltage-dependent blockade of sodium channels in a manner similar to anticonvulsants (Salgado, 1990, 1992; Deecher et al., 1991;

4 Sodium Channels

Deecher and Soderlund, 1991). The recent development of indoxacarb, the first commercial insecticide derived from pyrazoline-type compounds (Harder et al., 1996; Wing et al., 1998), underscores the continued relevance of the sodium channel as a target for insecticide development. Sodium channels are also the target of insect-selective polypeptide toxins from scorpion venoms (Zlotkin et al., 1995) which have been genetically engineered into insect baculoviruses; two potential biopesticide products based on this technology have proceeded to the stage of large-scale field tests (Cory, 1999). 5.1.1.3. Progress in Insect Sodium Channel Research Since 1985

The report in 1984 of the first DNA sequence for a voltage-sensitive ion channel, the voltage-sensitive sodium channel of the eel (Electrophorus electricus) electric organ (Noda et al., 1984), marked the beginning of a period of explosive growth in the application of molecular techniques to the study of ion channels that continues to the present. Since that time, studies of mammalian ion channels have identified the existence of gene families for both the a and b subunits of the voltage-sensitive sodium channel and have elucidated the structural domains of sodium channel a subunits that confer many of the biophysical and pharmacological properties of channel isoforms (Catterall, 2000; Goldin, 2001). Biomedical applications resulting from this research include the identification of sodium channel gene mutations that cause heritable diseases and a renewed interest in voltage-sensitive sodium channels as targets for therapeutic agents (Cannon, 2000; Clare et al., 2000; Baker and Wood, 2001; Goldin, 2001). The corresponding expansion of information on voltage-sensitive sodium channels in insects during this period has occurred in two stages. The first of these involved the integration of molecular biology with classical genetic approaches in Drosophila melanogaster to further illuminate the nature of gene mutations that were implicated genetically as determinants of sodium channel function and to determine the molecular structures of the genes defined by these mutations. The second phase, in which the molecular and functional characterization of sodium channels was extended to include numerous other insect taxa, was driven principally by research to define the role of sodium channel gene mutations in insecticide resistance. This chapter reviews the literature on insect voltage-sensitive sodium channels since 1985 from both of these perspectives.

5.1.2. Sodium Channel Genes in Insects 5.1.2.1. Sodium Channel Genes in Drosophila melanogaster

5.1.2.1.1. dsc1 The first putative insect sodium channel a subunit gene to be identified, dsc1, was isolated from D. melanogaster DNA libraries by low-stringency hybridization to a probe from the E. electricus sodium channel (Okamoto et al., 1987; Salkoff et al., 1987; Ramaswami and Tanouye, 1989). Sequence analysis of partial genomic DNA and cDNA clones identified sequences encoding the four internally homologous domains (Salkoff et al., 1987), but sequences of the 50 and 30 termini and the segments lying between homology domains I and II and homology domains II and III, all of which are regions of low sequence conservation in vertebrate genes, were not defined until the recent report of a full-length dsc1 coding sequence (Kulkarni et al., 2002). The dsc1 locus was initially mapped to cytogenetic region 60D-E on chromosome 2R (Okamoto et al., 1987; Salkoff et al., 1987; Ramaswami and Tanouye, 1989) and subsequently localized to cytogenetic region 60E5 by genome sequencing (Adams et al., 2000). Expression of the dsc1 gene is under both anatomical and developmental regulation: expression in embryos and larvae is restricted to very few cells, some of which may be nonneuronal, whereas dsc1-derived transcripts are found at much higher levels in pupae and adults, where they are widely expressed in the central nervous system and retina (Tseng-Crank et al., 1991; Hong and Ganetzky, 1994). A recent genome-wide screen identified dsc1 transcripts as targets for RNA editing at a site in the ‘‘inactivation gate’’ sequence (Figure 1) between homology domains III and IV (Hoopengardner et al., 2003). Alignment of the dsc1 protein sequence with those of vertebrate sodium channels suggests that dsc1 may exhibit atypical ion selectivity. Two conserved amino acid residues, a lysine in the pore-forming region of homology domain III and an alanine in the corresponding location in homology domain IV, are crucial for the sodium ion selectivity typical of voltage-sensitive sodium channels. When either of these is mutated to a glutamate residue the resulting channel becomes permeable to calcium and other monovalent and divalent cations that normally do not permeate sodium channels (Heinemann et al., 1992). The dsc1 protein sequence contains a glutamate residue rather than the conserved lysine residue in the pore-forming region of homology domain III and therefore may form a channel that is permeable to calcium as well as sodium. This observation is intriguing in light of recent studies identifying a

Sodium Channels 5

novel sodium current in dorsal unpaired median neurons of the cockroach Periplaneta americana that is biophysically distinct from the more typical sodium channels present in the same neurons and is permeable to calcium as well as sodium (Grolleau and Lapied, 2000; Defaix and Lapied, 2001). 5.1.2.1.2. para The parats (paralytic–temperaturesensitive) locus at cytogenetic location 14D on the X chromosome of D. melanogaster was first identified as the site of a temperature-sensitive paralytic mutation (Suzuki et al., 1971), which was subsequently found to cause temperature-sensitive impairment of action potential conduction in nerves (Siddiqi and Benzer, 1976; Wu and Ganetzky, 1980). Sequencing of genomic DNA and cDNA clones derived from the para locus showed that para encodes a protein having all of the structural hallmarks of voltage-sensitive sodium channel a subunits and exhibiting approximately 50% overall amino acid

sequence identity to vertebrate sodium channel a subunits (Loughney et al., 1989). The para gene product is quite divergent from the dsc1 gene product in that the four conserved internal homology domains of the two D. melanogaster genes are only as similar to each other (
50% amino acid sequence identity) as they are to the corresponding regions of vertebrate sodium channel proteins. Sequence analyses of partial cDNAs derived from the para loci of D. melanogaster and the related species D. virilis identified alternative mRNA splicing at eight sites involving seven optional exons (designated a, b, e, f, h, i, and j) and one pair of mutually exclusive exons (designated c/d), resulting in 256 possible unique structural variants from the para gene (Thackeray and Ganetzky, 1994, 1995; O’Dowd et al., 1995; Warmke et al., 1997). The locations of alternatively spliced exons in relation to other structural landmarks in the para protein are illustrated in Figure 2. Alternative splicing at

Figure 2 Diagram showing the approximate size and location of optional and mutually exclusive exons of in the para and Vssc1 sodium channel genes in relation to other sodium channel structural landmarks. (Modified with permission from Lee, S.H, Ingles, P.J., Knipple, D.C., Soderlund, D.M., 2002. Developmental regulation of alternative exon usage in the housefly Vssc1 sodium channel gene. Invert. Neurosci. 4, 125–133.)

6 Sodium Channels

these sites produces a heterogeneous family of sodium channel a subunit transcripts. Additional transcript heterogeneity in these species is conferred by posttranscriptional RNA editing at three sites (Hanrahan et al., 2000). Transcripts from the para locus are expressed strongly in the nervous system throughout development, but analysis of transcript pools from different developmental stages and anatomical regions revealed significant developmental and anatomical regulation of both alternative splicing and RNA editing (Tseng-Crank et al., 1991; Hong and Ganetzky, 1994; Thackeray and Ganetzky, 1994, 1995; Hanrahan et al., 2000). 5.1.2.1.3. tipE The tipE (temperature-induced paralysis, locus E) gene, at cytogenetic region 64B2, is the site of a temperature-sensitive paralytic mutation having a phenotype similar to that of para mutants (Kulkarni and Padhye, 1982; Jackson et al., 1986; Feng et al., 1995b). Sequence analysis of cDNAs derived from the tipE locus identified an open reading frame encoding a 452-amino acid protein that is clearly not a member of the voltagesensitive ion channel family and exhibits no significant sequence similarity to any other proteins in sequence databases (Feng et al., 1995a). The tipE protein contains two hydrophobic segments having properties consistent with the formation of transmembrane helices that flank a loop that contains five potential N-glycosylation sites. Transcripts from the original tipE mutant strain contain a premature stop codon between the two hydrophobic domains and therefore express a truncated tipE protein. Expression of the tipE gene in D. melanogaster is restricted to the nervous system and exhibits strong developmental regulation, so that transcript levels are very low in embryonic and larval stages but increase markedly in the late pupal stage. Genetic and functional criteria (see Section 5.1.3.1) suggest that the tipE protein may be a sodium channel auxiliary subunit, functionally analogous but structurally unrelated to vertebrate sodium channel b subunits. 5.1.2.1.4. Genetic and functional characterization of D. melanogaster mutants Gene dosage and interaction studies with parats mutants provide evidence for a crucial physiological role for the para protein. Homozygous parats mutants that exhibit temperature-sensitive paralysis in a wild-type background are unconditionally lethal in the presence of either napts (no action potential, temperaturesensitive) or tipE, both of which also impair nerve function at nonpermissive temperatures (Ganetzky, 1984, 1986). Similarly, decreased dosage of paraþ is unconditionally lethal in a napts background,

but increased dosage of paraþ causes leg-shaking behavior, a sign of neuronal excitation, and suppresses the temperature sensitivity of napts (Stern et al., 1990). The smellblind locus was originally identified as the site of mutations causing olfactory defects and was mapped close to para on the X chromosome (Lilly and Carlson, 1989). Two viable smellblind alleles exhibit temperature-dependent lethality to embryos at high temperatures and to adults at low temperatures, whereas four other alleles are recessive lethals (Lilly and Carlson, 1989; Lilly et al., 1994b). None of the six smellblind alleles complements the lethality of two unconditionally lethal para alleles, and two of the recessive lethal alleles of smellblind contain rearrangements within the primary para transcript (Lilly et al., 1994a). Taken together, these results show that smellblind is a novel class of para mutation. Interestingly, smellblind alleles do not exhibit the rapid, temperaturesensitive paralysis of parats mutants, and the parats mutants examined so far do not exhibit olfactory defects (Lilly et al., 1994a). Electrophysiological and pharmacological studies provide additional evidence for the functional importance of the para gene product. The temperaturesensitive failure of nerve conduction associated with parats alleles is presynaptic to the cervical giant fibers, which remain electrically excitable at nonpermissive temperatures (Nelson and Wyman, 1989; Elkins and Ganetzky, 1990). Cultured neurons from embryos homozygous for different parats alleles exhibit an allele-dependent reduction in sodium current density; in the case of paraST76 the expression of sodium currents was reduced by 98% (O’Dowd et al., 1989). One of these alleles, parats2, also caused a depolarizing shift in the voltage dependence of activation of sodium channels in cultured embryonic neurons. Cultured larval neurons from parats individuals exhibit temperaturedependent resistance to veratridine, which binds to site 2 on the sodium channel and causes persistent channel activation (Suzuki and Wu, 1984). Also, several parats strains are resistant to DDT and pyrethroid insecticides (Pittendrigh et al., 1997). The tipE mutation causes a temperaturedependent paralytic phenotype similar to that caused by parats alleles (Jackson et al., 1986). Three additional mutant tipE alleles, generated by g-ray mutagenesis and characterized as deletions or translocations disrupting the tipE locus, confer the temperature-sensitive phenotype when heterozygous with the original ethylmethanesulfonateinduced tipE allele (Feng et al., 1995b). This finding suggests that the original tipE mutation, identified

Sodium Channels 7

as a premature stop codon yielding a truncated gene product (Feng et al., 1995a), is a loss-of-function mutation. In addition to conferring the conditional paralytic phenotype, the tipE mutation causes temperature-sensitive impairment of conduction in adult, but not larval, nerves (Ganetzky, 1986; Elkins and Ganetzky, 1990) and reductions of approximately 50% in the sodium current density of cultured embryonic neurons and in density of [3H]saxitoxin binding sites in adult head preparations (Jackson et al., 1986). Studies using cultured embryonic neurons show that tipEþ is required for the sustained repetitive firing of sodium-dependent action potentials (Hodges et al., 2002). Sodium currents in neurons from animals carrying the tipE mutation recover more slowly, and the minimum interpulse interval needed to produce a second full-amplitude action potential is longer, than those in neurons from wildtype animals. Expression of the wild-type tipEþ transgene rescues all of the functional deficits conferred by the tipE mutation in these assays. Interactions between tipE and parats alleles provide further insight into the functional role of the tipE protein. As noted above, tipE interacts with various parats alleles to cause unconditional lethality at temperatures that are permissive for single mutants (Ganetzky, 1986). It is interesting that some alleles of para exhibit partial viability when combined with tipE, whereas other allelic combinations are unconditionally lethal. These observations have been interpreted as evidence that the tipE and para proteins may interact physically (Ganetzky, 1986). In contrast to the large body of information available about the functional significance of para and tipE, there is very little information on the role of dsc1. Embryonic neurons homozygous for a deficiency that includes the dsc1 locus exhibit normal sensitivity to veratridine (Sakai et al., 1989), thereby indicating the presence of functional sodium channels in these cells. Moreover, these neurons express voltage-sensitive sodium currents that are indistinguishable from those in wild-type cells (Germeraad et al., 1992). These studies provide further evidence that para rather than dsc1 encodes the sodium channels found in embryonic neurons. Until recently, there were no mutations associated with the dsc1 locus that might illuminate the function of the dsc1 protein. However, a series of smellimpaired (smi) mutants, created by single P element insertion mutagenesis and isolated on the basis of the loss of an olfactory avoidance behavioral phenotype, included one mutation (smi60E) mapping close to the dsc1 locus (Anholt et al., 1996). Subsequent studies showed that this mutant strain contains a P element transposon within the second

intron of the dsc1 gene (Kulkarni et al., 2002). The reduction of olfactory avoidance behavior in this strain is correlated with a reduction in the abundance of dsc1 transcripts, and excision of the P element restores wild-type behavior and dsc1 transcript levels. These results provide the first clear indication of a physiologically defined role for the dsc1 protein. 5.1.2.2. Sodium Channel Genes in Other Insect Species

5.1.2.2.1. Orthologs of para The identification of the para gene product as a physiologically important voltage-sensitive sodium channel in D. melanogaster (Loughney et al., 1989) and the concurrent development of the polymerase chain reaction (PCR) as a means of gene identification (Saiki et al., 1988) provided the conceptual and technical framework for the isolation of para-orthologous DNA sequences from other arthropod species (Doyle and Knipple, 1991; Knipple et al., 1991). The likely importance of voltage-sensitive sodium channels as sites of mutations conferring resistance to DDT and to pyrethroid insecticides provided further impetus for this effort (Soderlund and Knipple, 2003). As a result, full-length or partial genomic DNA or cDNA sequences now are known for para-orthologous sodium channel genes from at least 15 additional arthropod species. Two concurrent and independent efforts led to the characterization of the full-length coding sequence of Vssc1 (also called Msc), the para ortholog of the housefly (Musca domestica) (Ingles et al., 1996; Williamson et al., 1996). The inferred sequence of the Vssc1 protein is 90% identical to that of the most similar splice variant of the para gene product (Loughney et al., 1989; Thackeray and Ganetzky, 1994). The initial sequence analyses of Vssc1 cDNA clones obtained from adult fly heads did not identify alternatively spliced transcripts (Ingles et al., 1996; Williamson et al., 1996). A subsequent study investigated the issue of alternative splicing at the Vssc1 locus in greater detail by examining the heterogeneity of multiple cDNA clones derived from Vssc1 transcripts in three different developmental stages and two different adult body regions (Lee et al., 2002). This investigation identified multiple alternative exons in the Vssc1 gene, including a new pair of mutually exclusive exons (designated k and l) (Figure 2) encompassing part of transmembrane domain IIS3 and all of transmembrane domain IIS4. This splice site was not previously identified in the analysis of para transcripts in Drosophila species, which contain a segment homologous to exon l (Thackeray and Ganetzky, 1994, 1995; O’Dowd

8 Sodium Channels

et al., 1995; Warmke et al., 1997), but a search of the para genomic sequence (Adams et al., 2000) identified a segment upstream of the exon l-like sequence with substantial sequence similarity to exon k of Vssc1 (Lee et al., 2002). The amino acid sequences of exons k and l are highly divergent, differing at 16 of 41 amino acid residues, but retain conserved features of the voltage sensor structure in domain IIIS4. Other novel aspects of alternative splicing of Vssc1 transcripts in M. domestica included the apparent constitutive expression of optional exons h and i and the apparent inactivation of exon c by a stop codon, so that all transcripts encoding full-length channel sequences contained only exon d at this site. Alternative splicing at all of the sites identified either in Vssc1 or in the para genes of D. melanogaster or D. virilis could theoretically generate up to 512 structurally unique sodium channel splice variants. However, only a small subset of these variants (five or fewer) comprised more than 90% of the transcripts in each M. domestica cDNA pool examined. The para ortholog of Blattella germanica (designated paraCSMA) encodes an inferred protein sequence that is 76–78% identical to the two dipteran sequences (Dong, 1997; Tan et al., 2002a). Analysis of multiple paraCSMA sequences identified novel aspects of alternative exon usage at some of the sites of alternative splicing found in D. melanogaster and M. domestica. For example, paraCSMA transcripts contain two examples of segments corresponding to optional exon j, suggesting that the B. germanica gene may incorporate multiple variants of this exon. Similarly, the B. germanica transcripts contain three different exons (designated G1, G2, and G3) at the exon k/l splice site in homology domain III (Tan et al., 2002a). Exons G1 and G2 correspond to exons l and k of para and Vssc1, whereas exon G3 exhibits no obvious sequence similarity to either exon G1 or G2 and contains an in-frame stop codon. Exon G3 corresponds exactly to an alternative exon in vertebrate Nav1.6 sodium channels that contains a premature stop codon at the same position (Plummer et al., 1997). The structure of hscp, the ortholog of para in Heliothis virescens, was determined by sequence analyses of genomic DNA and PCR-amplified cDNA segments (Park et al., 1999). The reported hscp coding sequence, which is incomplete at the 30 end, gives a predicted protein sequence that is 80% identical to the corresponding coding region of para. Fifteen of the 19 introns identified in the hscp gene were conserved in location with respect to the corresponding introns in para, and alternatively spliced transcripts containing optional exon j and mutually exclusive exons c and d were detected. This study also

identified several transcripts lacking sequences from the linker between homology domains I and II that contained premature stop codons. Unlike the truncation of the M. domestica and B. germanica transcripts by premature stop codons located in alternatively spliced exons, these H. virescens transcripts appear to be the result of splicing errors that resulted in missing exons and frameshifts. The ortholog of para in the head louse, Pediculus capitis, encodes a voltage-sensitive sodium channel a subunit protein that is 73% identical to para and the B. germanica and M. domestica orthologs (Lee et al., 2003). Full-length sequences from two strains of head louse and one strain of body louse (P. humanus) were 99.6% identical at the nucleotide level, providing further evidence that the head louse and body louse are conspecific. Analysis of multiple transcripts provided evidence for alternative splicing of a segment homologous to exon j of para. All of the transcripts contained segments corresponding to para optional exons a, b, and h and mutually exclusive exons d and l but did not contain segments corresponding to optional exons i, e, and f. The ortholog of para in the varroa mite, Varroa destructor (designated VmNa) encodes a voltagesensitive sodium channel a subunit that is only 51% identical to the para gene product of D. melanogaster (Wang et al., 2003). Sequence analysis of multiple VmNa transcripts identified several novel aspects of alternative exon usage in this gene. Some VmNa transcripts contained an optional exon (designated exon 3) corresponding to the pair of mutually exclusive introns k and l in Drosophila species and M. domestica (Figure 2). Some VmNa transcripts also contained option exons at three sites not previously identified as sites of alternative splicing: exon 1, a segment of homology domain II containing part of the IIS2 and all of the IIS3 transmembrane helices; exon 2, a short inserted sequence in the domains II–III intracellular linker; and a retained intron in the C terminus containing an alternative stop codon. Channels lacking exon 1 or 3 would also lack critical determinants of channel organization and function and therefore may be inactive, whereas the alternative splicing of exon 2 and the alternative C terminus could be involved in the regulation of channel expression or functional properties. Partial sequences of varying length also exist for the para orthologs from 10 additional arthropod species: Anopheles gambiae (Martinez-Torres et al., 1998; Ranson et al., 2000); Bemisia tabaci (Morin et al., 2002); Boophilus microplus (He et al., 1999); Culex pipiens (Martinez-Torres et al., 1999a); Frankliniella occidentalis (Forcioli et al., 2002); Helicoverpa armigera (Head et al., 1998); Hematobia

Sodium Channels 9

irritans (Guerrero et al., 1997); Leptinotarsa decemlineata (Lee et al., 1999b); Myzus persicae (MartinezTorres et al., 1999b); and Plutella xylostella (Schuler et al., 1998).

p0145

5.1.2.2.2. Orthologs of dsc1 The search for the para ortholog of B. germanica also yielded a partial cDNA of the ortholog of dsc1 in this species (Dong, 1997). Subsequent work described the complete coding sequence of this gene (designated BSC1) and identified three regions of alternative exon usage (Liu et al., 2001). Reverse transcription (RT)-PCR analyses documented the expression of BSC1 transcripts not only in the nerve cord but also in muscle, gut, fat body, and ovary, a much broader pattern of expression than was found by in situ hybridization of dsc1-derived probes to D. melanogaster tissues (Hong and Ganetzky, 1994). RT-PCR assays also showed that the expression of BSC1 splice variants was both tissue-specific and developmentally regulated (Liu et al., 2001). So far, the only other ortholog of dsc1 described in the literature is a partial genomic DNA sequence from H. virescens (Park et al., 1999). 5.1.2.2.3. Orthologs of tipE The tipE gene is of interest because of its clear involvement in modifying sodium channel expression and function and the suggestion that it may encode a novel type of sodium channel auxiliary subunit. The ortholog of tipE in Musca domestica, designated Vsscb, encodes a predicted protein that exhibits 72% overall amino acid sequence identity to the tipE protein and 97% identity within the hydrophobic regions identified as probable transmembrane domains (Lee et al., 2000a). These results suggest that tipE and Vsscb are substantially more divergent in sequence than are the two sodium channel a subunit genes from these species.

5.1.3. Functional Expression of Cloned Insect Sodium Channels 5.1.3.1. Expression of Functional Sodium Channels in Xenopus Oocytes

Unfertilized oocytes of the African clawed frog, Xenopus laevis, are a powerful tool for confirming the functional roles of neurotransmitter receptors and ion channels and for correlating channel structure with functional and pharmacological properties (Lester, 1988). Oocytes injected with cRNA (synthetic mRNA prepared from cloned cDNA) express sodium channels in the cell membrane that can be detected by conventional electrophysiological

assays, such as two-electrode voltage clamp analysis of the currents carried by the expressed proteins in response to changes in membrane potential or the application of native or exogenous ligands (Goldin, 1992; Stu¨ hmer, 1992). This system, coupled with site-directed mutagenesis of cloned cDNAs, has been widely employed in structure–function studies of vertebrate sodium channels (Catterall, 2000; Goldin, 2001; Yu and Catterall, 2003). Initial efforts to express functional sodium channel from synthetic para cRNA injected into oocytes produced voltage-gated sodium currents of very low amplitude, but coexpression of para and tipE cRNAs stimulated sodium current expression approximately 30-fold (Feng et al., 1995a). Subsequent studies (Warmke et al., 1997) showed that injection of large amounts of para cRNA was required to obtain more robust currents in the absence of tipE cRNA. These authors also showed that para/tipE channels inactivated more rapidly than those expressed from para alone. Thus, coexpression of para and tipE reconstitutes the functional properties of the native D. melanogaster voltage-sensitive sodium channel and provides evidence that the tipE protein may function as a sodium channel auxiliary subunit. Efforts to express functional sodium channels in oocytes from Vssc1 cRNA produced results similar to those with para, requiring large amounts of injected cRNA to yield low levels of sodium current expression, but coexpression with tipE cRNA resulted in the robust expression of voltage-gated sodium currents (Smith et al., 1997; Lee et al., 2000a). These Vssc1/tipE channels, like para/tipE channels, inactivated more rapidly than channels expressed from either para or Vssc1 cRNA alone. Coexpression of Vssc1 with its conspecific auxiliary subunit, Vsscb, was more effective than coexpression with tipE in enhancing the level of sodium current expression and accelerating the inactivation kinetics of Vssc1 sodium channels (Lee et al., 2000a). Coexpression with tipE cRNA was also found to be necessary for the expression of functional sodium channels from paraCSMA cRNA in oocytes (Tan et al., 2002b). However, this study did not document the expression of functional channels in oocytes from paraCSMA cRNA alone. 5.1.3.2. Pharmacological Properties of Expressed Insect Sodium Channels

Insect sodium channels expressed in oocytes retain sensitivity to insecticides and other natural and synthetic toxicants that alter sodium channel function. Insect sodium channels expressed in oocytes also are sensitive to blockade by nanomolar concentrations of tetrodotoxin (Warmke et al., 1997; Smith et al.,

10 Sodium Channels

1998; Tan et al., 2002a). These channels are also susceptible to modification by the alkaloid batrachotoxin (Lee and Soderlund, 2001) and polypeptide toxins from sea anemone (Warmke et al., 1997) and scorpion (Shichor et al., 2002) venoms. The effects of pyrethroids on expressed insect sodium channels have been extensively investigated in the context of the functional characterization of sodium channel mutations associated with pyrethroid resistance (see Section 5.1.4.5). When assayed under voltage clamp conditions, type I pyrethroids (i.e., cismethrin and permethrin) appear to bind predominantly to resting or inactivated channels, shifting the voltage dependence of activation to more negative potentials and causing a slowly activating sodium current. These compounds also produce characteristic sodium tail currents following a depolarizing pulse that decay with first-order time constants (Smith et al., 1997, 1998; Warmke et al., 1997; Zhao et al., 2000). In contrast to these results, type II pyrethroids (i.e., [1R,cis,aS]-cypermethrin and deltamethrin) exhibit profound use-dependent modification of sodium currents, which implies that these compounds bind preferentially to activated sodium channel states (Smith et al., 1998; Vais et al., 2000; Tan et al., 2002b). The tail currents caused by these compounds are more persistent than those caused by cismethrin or permethrin. In the case of deltamethrin, tail current decay is biphasic, rather than monophasic (Vais et al., 2000; Tan et al., 2002b); this finding has been interpreted as evidence for two binding sites on insect sodium channels with different affinities for this ligand (Vais et al., 2000). 5.1.3.3. Functional Characterization of Sodium Channel Splice Variants

The conservation of alternative exon structure and the developmental and anatomical regulation of alternative exon usage in the para sodium channel gene of D. melanogaster and its orthologs in other insect species imply that alternative splicing may generate a family of sodium channel proteins with differing functional properties, as has been found for other ion channels and receptors (HarrisWarwick, 2000). Most of the optional exons identified in para orthologs to date (see Figure 2) are located in intracellular domains of the sodium channel protein. These exons may therefore be involved in the regulation of sodium channel expression or function as the result of interactions with protein kinases or G proteins (Cukierman, 1996; Cantrell and Catterall, 2001). This interpretation is consistent with the existence in exons a and i of consensus protein kinase A phosphorylation sites (O’Dowd et al., 1995; Ingles et al., 1996). Similarly, exons

2 and 4 of the VmNa sodium channel are located in intracellular domains, and exon 2 contains contains a consensus protein kinase C phosphorylation site (Wang et al., 2003). The sole exception so far to the intracellular location of optional exons is exon 1 of the VmNa sodium channel sequence, which is located in the transmembrane region of domain II (Wang et al., 2003). So far, there is little direct evidence bearing on the functional significance of the alternative splicing of optional exons. In embryonic D. melanogaster neurons, functional sodium channels were detected only in those cells having para transcripts containing exon a (O’Dowd et al., 1995). This study also documented enhanced sodium current expression in cells expressing channels that contain both exons a and i. Whereas these findings imply a critical role for exon a alone and the combination of exons a and i together in sodium channel regulation, the direct comparison in functional expression assays using X. laevis oocytes of variants of para that differ only by the presence or absence of exon a did not find any effects of exon a on sodium current expression or properties in this system (Warmke et al., 1997). In contrast to the optional exons, the mutually exclusive exons in para orthologs occur within the transmembrane regions of homology domains II and III (Figure 2). There is no information on the functional role of the alternative splicing of exons c and d. In D. melanogaster, these exons differ by only two of 55 amino acid residues (Loughney et al., 1989). In M. domestica, all functional channels apparently contain only exon d because exon c contains an inframe stop codon (Lee et al., 2002). These observations suggest that alternative splicing at the c/d site may play a role in posttranscriptional regulation rather than in the generation of functionally distinct channel variants. The most significant functional effects of alternative exon usage have been documented for splice variants at the exon k/l site. Unlike exons c and d, exons k and l (corresponding to exons G2 and G1 in B. germanica) differ substantially in amino acid sequence (Lee et al., 2002; Tan et al., 2002a). Expression of paraCSMA variants containing either exon G1 or G2 in oocytes documents differences in the voltage dependence of both activation and inactivation of these channels (Tan et al., 2002a). Unexpectedly, this study also found substantial differences between these variants in their sensitivity to the pyrethroid insecticide deltamethrin. These results provide the first experimental evidence for functional differences between splice variants of insect sodium channels. Alternative splicing at the k/l site in

Sodium Channels 11

B. germanica and V. destructor also appears to be involved in the expression of inactive channel variants, in that exon G3 in the paraCSMA sequence encodes a truncated channel (Tan et al., 2002a) and the absence of exon 3 in the VmNa sequence encodes a channel lacking one of the four voltage sensor regions (Wang et al., 2003).

5.1.4. Sodium Channels and Knockdown Resistance to Pyrethroids 5.1.4.1. Knockdown Resistance

p0205

The knockdown resistance (kdr) trait, which confers resistance to the rapid knockdown action and lethal effects of DDT and pyrethrins, was first documented in houseflies in 1951 (Busvine, 1951) and isolated genetically in 1954 (Milani, 1954). The kdr trait confers resistance to both the rapid paralytic and lethal actions of all known pyrethroids, as well as the pyrethrins and DDT, but does not diminish the efficacy of other insecticide classes (Oppenoorth, 1985). Electrophysiological assays employing a variety of nerve preparations from larval and adult kdr insects (Bloomquist, 1988) provide direct evidence for reduced neuronal sensitivity as the basis for the kdr trait. A second resistance trait in the housefly (designated super-kdr) that confers much greater resistance to DDT and pyrethroids than that found in kdr strains has also been isolated genetically and mapped to chromosome 3 (Sawicki, 1978; Farnham et al., 1987). The kdr and super-kdr traits are widely presumed to represent allelic variants at a single resistance locus on the basis of their similar spectra of resistance and their common localization to chromosome 3 in the housefly. Resistance mechanisms similar to kdr have been inferred in a number of agricultural pests and disease vectors on the basis of cross-resistance patterns and the absence of synergism by compounds known to inhibit the esterase and monooxygenase activities involved in pyrethroid metabolism (Soderlund and Bloomquist, 1990; Bloomquist, 1993; Soderlund, 1997; Soderlund and Knipple, 1999). Confirming electrophysiological evidence for reduced neuronal sensitivity to pyrethroids also exists for at least six species: H. virescens, Spodoptera littoralis, Culex quinquefasciatus, A. stephensi, B. germanica, and P. xylostella (Bloomquist, 1988, 1993; Schuler et al., 1998). Pyrethroids are known to exert their insecticidal effects by altering the function of voltage-sensitive sodium channels in nerve membranes (see Section 5.1.1.2). Therefore, studies of kdr-like resistance have focused on mechanisms that might affect the

regulation, pharmacology, or function of the sodium channel. 5.1.4.2. Altered Sodium Channel Regulation as a Mechanism of Knockdown Resistance

The possible role of reduced insecticide receptor density in knockdown resistance was initially implicated on the basis of resistance-associated reductions in the density of binding sites for [3H]STX (Rossignol, 1988), a radioligand that specifically labels site 1 of the sodium channel (see Table 1). However, further investigation of [3H]STX binding in susceptible, kdr, and super-kdr housefly strains (Grubs et al., 1988; Sattelle et al., 1988; Pauron et al., 1989) revealed that a reduction in sodium channel density is not an obligatory component of the kdr or super-kdr phenotypes of the housefly. Moreover, comparisons of Vssc1 transcript and protein levels in susceptible and kdr housefly strains did not document any differences between strains (Castella et al., 1997). Although these results rule out sodium channel downregulation as the mechanism underlying the kdr and super-kdr traits in the housefly, reduced sodium channel density could, in principle, produce a kdr-like phenotype. The relationship between sodium channel density and kdr-like resistance has been evaluated directly using the napts strain of D. melanogaster, in which the density of sodium channels (measured as binding sites for [3H]STX in head membrane preparations) is approximately half that of wild-type flies (Jackson et al., 1984). Flies homozygous for napts exhibit modest (
threefold) resistance to the lethal effects of DDT and pyrethroids that is also evident in delayed onset of paralysis and reduced physiological sensitivity of the adult central nervous system (Kasbekar and Hall, 1988; Bloomquist et al., 1989). These results suggest that if such a mechanism were present in other insects that exhibit knockdown resistance, the magnitude of resistance observed would require a profound and readily detectable reduction in sodium channel density (Grubs et al., 1988). Because reductions in sodium channel density more severe than that observed in the napts strain of D. melanogaster would be anticipated to compromise viability, it is unlikely that reduced target density can account for kdr-like traits that confer significant levels of resistance. 5.1.4.3. Genetic Linkage between Knockdown Resistance and Sodium Channel Genes

Two studies employed restriction fragment length polymorphisms (RFLPs) in the Vssc1 gene coupled with discriminating dose bioassays with DDT to demonstrate tight genetic linkage (within
1 map

12 Sodium Channels

p0230

unit) of the kdr and super-kdr resistance trait and the Vssc1 gene of M. domestica (Williamson et al., 1993; Knipple et al., 1994). In addition to providing strong genetic evidence for mutations at a sodium channel structural gene as the cause of knockdown resistance in the housefly, these two studies also provided the first experimental evidence for the widely presumed allelism of the kdr and super-kdr traits in this species. Conceptually similar approaches were employed to investigate linkage between knockdown resistance traits and para-orthologous sodium channel sequences in other species. In the case of B. germanica, knockdown resistance was tightly linked (within
0.2 map units) to an RFLP located with the para-orthologous sodium channel gene (Dong and Scott, 1994). The use of RFLP markers to assess the linkage between knockdown resistance and sodium channel gene sequences in H. virescens (Taylor et al., 1993) was complicated by the use of a strain with multiple resistance mechanisms. Nevertheless, results of these assays suggested that one component of resistance was linked to an RFLP in the para-orthologous sodium channel gene of this species. In L. decemlineata (Lee et al., 1999b) and B. tabaci (Morin et al., 2002), sequencing of DNA

from individual insects of susceptible and resistant phenotypes has identified sequence variants within sodium channel coding regions that are genetically linked with resistant phenotypes. Finally, some mutant alleles of para that exhibit the temperaturesensitive paralytic phenotype also exhibit resistance to pyrethroids at permissive temperatures (Hall and Kasbekar, 1989; Pittendrigh et al., 1997). 5.1.4.4. Identification of Resistance-Associated Mutations

The selective alteration of sodium channel pharmacology in knockdown resistant insects and the genetic linkage of knockdown resistance traits and sodium channel gene sequences provided a strong impetus for the identification and functional characterization resistance-associated mutations in insect sodium channel genes. Results of these efforts are the subject of two recent comprehensive reviews (Soderlund and Knipple, 1999, 2003). Comparison of partial and complete sequences from 15 housefly strains representing multiple examples of susceptible, kdr, and super-kdr phenotypes consistently identified two point mutations that were associated with resistant phenotypes (Figure 3): mutation of leucine to phenylalanine at

Figure 3 Locations and identities of sodium channel point mutations associated with knockdown resistance to pyrethroids. Symbols indicate the species in which each mutation was first identified (see also Table 2). (Modified with permission from Soderlund, D.M., Knipple, D.C. 2003. The molecular biology of knockdown resistance to pyrethroid insecticides. Insect Biochem. Mol. Biol. 33, 563–577.)

p0240

Sodium Channels 13

amino acid residue 1014 (designated L1014F) in all kdr and super-kdr strains, and the additional mutation of methionine to threonine at residue 918 (designated M918T) only in super-kdr strains (Ingles et al., 1996; Miyazaki et al., 1996; Williamson et al., 1996). Mutations in para-orthologous sodium channel gene sequences corresponding to the L1014F mutation in the housefly have been also identified to date in eight additional pest species (Table 2): B. germanica (Miyazaki et al., 1996; Dong, 1997), F. occidentalis (Forcioli et al., 2002), H. irritans (Guerrero et al., 1997), A. gambiae (MartinezTorres et al., 1998), P. xylostella (Schuler et al., 1998), L. decemlineata (Lee et al., 1999b), M. persicae (Martinez-Torres et al., 1999b), and C. pipiens (Martinez-Torres et al., 1999a). (For clarity and consistency and to facilitate comparisons between species, including those for which full-length sodium channel sequences are not available, all resistanceassociated mutations in this report are numbered according to their positions in the amino acid sequence of the most abundant splice variant of the housefly Vssc1 sodium channel a subunit (GenBank

t0010

Accession no. U38813).) In contrast to the many examples of mutations in other species corresponding to the L1014F mutation in the housefly, a mutation corresponding to the second-site M918T mutation that is associated with the super-kdr trait of the housefly has been found to date only in highly resistant populations of H. irritans (Guerrero et al., 1997). The search for sodium channel gene mutations associated with knockdown resistance has also identified numerous novel mutations (Figure 3; Table 2). Studies with pyrethroid-resistant H. virescens populations identified a second mutation at sequence position 1014, L1014H (Park and Taylor, 1997), as well as three new mutations: V410M (Park et al., 1997), D1549V, and E1553G (Head et al., 1998). The latter two mutations were found together in sodium channel sequences from pyrethroid-resistant strains of both H. virescens and H. armigera (Head et al., 1998). Studies with knockdown-resistant populations of C. pipiens and A. gambiae identified a third variant at position 1014, L1014S (Martinez-Torres et al., 1999a;

Table 2 Sodium channel amino acid sequence polymorphisms associated with knockdown resistance in arthropod species Species

Mutations identified a

Reference

Anopheles gambiae

L1014F L1014S M918V; L925I L1014Fb L1014FþE435KþC785Rb L1014FþD59GþE435KþC785RþP1999Lb F1538I L1014F; L1014S I253N; A1410V; A1494V; M1524I L1014FþT929C D1549VþE1533G L1014H V410M D1549VþE1533G L1014FþM918T L1014F L1014F; L1014FþM918T

Martinez-Torres et al. (1998) Ranson et al. (2000) Morin et al. (2002) Miyazaki et al. (1996), Dong (1997) Liu et al. (2000) Liu et al. (2000) He et al. (1999) Martinez-Torres et al. (1999) Pittendrigh et al. (1997) Forcioli et al. (2002) Head et al. (1998) Park and Taylor (1997) Park et al. (1997) Head et al. (1998) Guerrero et al. (1997) Lee et al. (1999b) Ingles et al. (1996), Miyazaki et al. (1996), Williamson et al. (1996) Martinez-Torres et al. (1999b) Lee et al. (2000b), (2003) Schuler et al. (1998) Wang et al. (2002)

Bemisia tabaci Blattella germanica

Boophilus microplus Culex pipiens Drosophila melanogaster Frankliniella occidentalis Helicoverpa armigera Heliothis virescens

Hematobia irritans Leptinotarsa decemlineata Musca domestica Myzus persicae Pediculus capitis Plutella xylostella Varroa destructor

L1014F M827IþT929IþL932Fc L1014FþT929I L1596PþM1823I F1528LþL1596PþI1742VþM1823I

a Positions numbered according to the amino acid sequence of the most abundant splice variant of the housefly Vssc1 sodium channel protein (Ingles et al., 1996; Williamson et al., 1996). b These mutations correspond to the D58G, E434K, C764R, L993F, and P1880L mutations in the full-length Blattella germanica paraCSMA sodium channel (Dong, 1997; Liu et al., 2000). c These mutations correspond to the M815I, T917I, and L920F mutations in the full-length Pediculus capitis sodium channel (Lee et al., 2003). Modified from Soderlund, D.M., Knipple, D.C. 2003. The molecular biology of knockdown resistance to pyrethroid insecticides. Insect Biochem. Mol. Biol. 33, 563–577.

p0245

14 Sodium Channels

p0250

p0255

Ranson et al., 2000). A novel mutation at Met918 (M918V) was recently identified in some pyrethroidresistant B. tabaci populations (Morin et al., 2002). Characterization of sodium channel sequences from pyrethroid-resistant P. xylostella identified a novel putative second-site mutation (T929I) associated with the more commonly observed L1014F mutation in strains with high pyrethroid resistance (Schuler et al., 1998). The T929I mutation is also observed in combination with two other novel mutations, M827I and L932F, in pyrethroid-resistant P. capitis (Lee et al., 2000b, 2003). A second mutation at position 929 (T929C) was found in combination with the L1014F mutation in highly resistant populations of F. occidentalis (Forcioli et al., 2002). Recent studies with B. tabaci have also identified another novel mutation in this region of the sodium channel protein (L925I) that is tighly linked to pyrethroid resistance (Morin et al., 2002). Four novel putative second-site mutations (D59G, E435K, C785R, and P1999L) are found together with the L1014F mutation in one or more strains of B. germanica that exhibit high levels of pyrethroid resistance (Liu et al., 2000). Finally, a screen for pyrethroid resistance in strains of D. melanogaster having temperature-sensitive paralytic phenotypes that map to the para sodium channel identified four novel resistance-associated mutations: I253N, A1410V, A1494V, and M1524I (Pittendrigh et al., 1997). The identification of mutations associated with pyrethroid resistance in para-orthologous sodium channel gene sequences also extends to noninsect arthropod species. Sodium channel sequences from populations of B. microplus that exhibit very high levels of pyrethroid resistance contain the F1538I mutation (He et al., 1999). Also, the para-orthologous sodium channel sequences obtained from populations of the mite V. destructor that are resistant to the pyrethroid fluvalinate contain four novel mutations (F1528L, P1596L, I1742V, and V1823I) (Wang et al., 2002), which include the first resistance-associated mutations identified in homology domain IV (Figure 3). Among the 26 unique sodium channel amino acid sequence polymorphisms associated so far with pyrethroid resistance, those occurring at five sites have been found as single mutations in resistant populations: Val410 (V410M in H. virescens), Met918 (M918V in Bemisia tabaci); Leu925 (L925I in B. tabaci), Leu1014 (L1014F in several species, L1014H in H. virescens, and L1014S in C. pipiens and A. gambiae); and Phe1538 (F1538I in B. microplus). Mutations at six sites (M918T in M. domestica and H. irritans; T929I in P. xylostella

and T929C in F. occidentalis; D59G, E435K, C785R, and P1999L in B. germanica) have been found in combination with the L1014F mutation in highly resistant strains and therefore have been hypothesized to function as second-site mutations that produce additive or synergistic enhancement the resistance caused by the L1014F mutation. The functional status of the remaining resistance-associated mutations is more ambiguous. 5.1.4.5. Functional Analysis of Resistance-Associated Mutations

The X. laevis oocyte expression system has been employed to characterize the effects of several resistance-associated mutations on the pyrethroid sensitivity and functional properties of expressed sodium channels. Most of these efforts have focussed on the mutations identified in sodium channels of kdr and super-kdr houseflies, but a more limited group of studies have also examined additional putative primary resistance mutations and certain putative secondary mutations. All of these studies involve the insertion of candidate mutations into wild-type sodium channel cDNAs, expression of wild-type and specifically mutated channels in oocytes, and direct comparison of the pharmacological properties of the expressed channels. 5.1.4.5.1. Functional analysis of the kdr and superkdr mutations The effects of the L1014F mutation, the sodium channel gene mutation most commonly associated with knockdown resistance, on pyrethroid sensitivity have been examined in several sodium channel sequence contexts and with different pyrethroids as probes. Housefly Vssc1 sodium channels containing the L1014F substitution, coexpressed in oocytes with the D. melanogaster tipE protein, were approximately tenfold less sensitive to modification by cismethrin, a type I pyrethroid, than wild-type Vssc1/tipE channels (Smith et al., 1997). The L1014F mutation also accelerated the rate of decay of cismethrin-induced sodium tail currents. Similar experiments with wild type and specifically mutated D. melanogaster para sodium channels coexpressed with the tipE protein and deltamethrin as the test pyrethroid confirmed and extended these findings (Vais et al., 2000). Comparison of the actions of deltamethrin on wild-type para/ tipE channels and channels containing the L1014F mutation reduced the sensitivity of expressed channels to deltamethrin approximately 17-fold and accelerated the rate of deltamethrin-induced tail current decay. More detailed analysis showed that mutated channels exhibited lower affinity for deltamethrin as well as a reduced availability of open

Sodium Channels 15

channel states due to enhanced closed-state inactivation. The effects of the L1014F mutation on deltamethrin sensitivity were also examined using B. germanica paraCSMA sodium channels coexpressed with tipE (Tan et al., 2002b). In this study, the L1014F mutation conferred approximately sixfold resistance to deltamethrin but, unlike the results obtained with Vssc1/tipE and para/tipE channels, the reduction in pyrethroid sensitivity of mutated paraCSMA/tipE channels was not accompanied by an acceleration of tail current decay. The discovery that the rat Nav1.8 (also called SNS or PN3) sodium channel isoform was highly sensitive to both type I and type II pyrethroids (Smith and Soderlund, 2001) provided the opportunity to examine the impact of the L1014F mutation in a sodium channel sequence environment that is otherwise substantially divergent (i.e., only
40% identical at the amino acid sequence level) from para-orthologous channels of insects. The L1014F mutation, introduced at the cognate conserved leucine residue of the rat Nav1.8 channel, reduced the sensitivity of expressed channels to cismethrin more than tenfold and increased the rate of decay of the cismethrin-induced tail current. These effects were qualitatively identical to the effects of the L1014F mutation on the cismethrin sensitivity of Vssc1/tipE channels (Smith et al., 1997). The effects of the M918T mutation in the presence of the L1014F mutation, the combination of mutations associated with the super-kdr resistance trait of the housefly, have also been examined in both insect and mammalian sodium channel sequence contexts with both type I and type II pyrethroids as probes. Incorporation of the M918T and L1014F mutations into the Vssc1 protein gave rise to Vssc1/tipE channels that were completely insensitive to both cismethrin and [1R,cis,aS]-cypermethrin at the highest concentrations that could be achieved given the low aqueous solubility of these compounds (Lee et al., 1999c). Similarly, rat Nav1.8 sodium channels containing the M918T/L1014F double mutation were completely insensitive to modification by the highest attainable concentration of cismethrin (Soderlund and Lee, 2001). In the para/tipE sequence context the M918T/L1014F double mutation decreased the sensitivity of expressed channels to deltamethrin approximately 100-fold and also produced tail currents with monophasic, rather than biphasic, decay kinetics (Vais et al., 2000). The latter effect was interpreted as evidence that the double mutation reduced the number of deltamethrin binding sites per channel from two to one. The effects of the M918T single mutation on pyrethroid sensitivity has also been evaluated in

multiple sequence contexts. Insertion of the M918T mutation into para/tipE channels gave channels that were twofold more resistant to deltamethrin than the doubly mutated (M918T/L1014F) channels; the M918T channels also exhibited monophasic tail current decay kinetics (Vais et al., 2001). Insertion of the M918T mutation in Vssc1/tipE sodium channels significantly impaired sodium current expression in oocytes and produced channels that were not detectably modified by cismethrin (Lee et al., 1999c). In rat Nav1.8 channels, the M918T mutation caused a degree of resistance to cismethrin that was equivalent to that caused by the single L1014F mutation (Soderlund and Lee, 2001). These results suggest that the M918T mutation does not enhance resistance caused by the L1014F mutation in an additive or synergistic manner but rather provides a high level of resistance that supercedes the effect of the L1014F mutation alone. In light of the profound reduction in pyrethroid sensitivity caused by the M918T single mutation in these assays, it is surprising that it has not been identified as a single mutation in pyrethroid-resistant insect populations. A likely explanation for this situation has emerged from studies of alternative exon usage in para and its orthologs (see Sections 5.1.2.1.2 and 5.1.2.2.1). In the para sequence of D. melanogaster, the Met918 residue occurs in mutually exclusive exons c and d, each of which is incorporated into a subpopulation of sodium channel sequence variants (Thackeray and Ganetzky, 1994). Exons homologous to exons c and d of D. melanogaster are also found among transcripts from the para-orthologous sodium channel gene of H. virescens (Park et al., 1999) and C. pipiens (Martinez-Torres et al., 1999a). In these situations, two independent point mutations (one each in exons c and d) would be required to insure that all channel variants contained the M918T substitution. In contrast, exon c in the housefly Vssc1 gene contains a stop codon, so that all full-length Vssc1 transcripts are derived solely from exon d (Lee et al., 2002). Thus, a single point mutation in exon d of the housefly is sufficient to insure that all functional sodium channel splice variants in this species contain the M918T substitution. This interpretation suggests that the M918T mutation and other mutations at this site (such as the M918V mutation in B. tabaci) can only be selected in species in which alternative splicing at the exon c/d locus produces functional channels from only one of these alternative exons. Although consistent with available data on alternative splicing at the Vssc1 locus in the housefly, this analysis does not explain the absence of the single M918T mutation in resistant strains of

p0285

16 Sodium Channels

this species. It therefore remains possible that a functional deficit associated with the M918T mutation is somehow complemented or rescued by the presence of the L1014F mutation (Lee et al., 1999c). 5.1.4.5.2. Functional analysis of other putative primary resistance mutations The effects of the V410M mutation, a single mutation associated with knockdown resistance in some H. virescens populations, has been examined in para/tipE, Vssc1/tipE, and paraCSMA/tipE sodium channels expressed in oocytes. Insertion of the V410M mutation into para/tipE channels resulted in a >10-fold reduction in permethrin sensitivity coupled with a
45-fold acceleration in the rate of sodium tail current decay (Zhao et al., 2000). Similar results were obtained in Vssc1/tipE channels containing the V410M mutation, which decreased channel sensitivity to cismethrin by
20-fold and accelerated the rate of tail current decay
10-fold (Lee and Soderlund, 2001). In paraCSMA/tipE channels, the V410M mutation caused
17-fold resistance to deltamethrin (Liu et al., 2002). In contrast to the effects of this mutation on the kinetics of channel modification by permethrin and cismethrin, resistance to deltamethrin in paraCSMA/tipE channels containing the V410M mutation was not accompanied by a significant acceleration in tail current decay kinetics. Although Met410 occurs in a region of the sodium channel known to be involved in the binding and action of batrachotoxin, the V410M mutation did not affect the potency of batrachotoxin as a modifier of Vssc1/tipE sodium channels in oocytes (Lee and Soderlund, 2001). The V410M mutation is the only resistance mutation that has been examined both in the oocyte expression system and in its native cellular context. Assays with cultured adult central neurons from H. virescens homozygous for the V410M mutation documented a
21-fold reduction in permethrin sensitivity of sodium channels in the resistant strain (Lee et al., 1999a). Although these authors did not report the time constants for the decay of permethrin-induced sodium tail current, inspection of their data suggests that reduced permethrin sensitivity in cultured neurons was apparently not correlated with a significant acceleration of tail current decay as was observed in some of the studies of the V410M mutation in oocytes (Zhao et al., 2000; Lee and Soderlund, 2001). In cultured neurons, channels containing the V410M mutation were approximately 2.6-fold more sensitive to the polypeptide scorpion toxin LqhaIT (Lee et al., 1999a). The characterization of mutations at Leu1014 was expanded to include the L1014H mutation,

another single mutation found in some knockdownresistant H. virescens populations (Zhao et al., 2000). Incorporation of the L1014H mutation into para/tipE sodium channels conferred >tenfold resistance to permethrin, a type I pyrethroid. Permethrin-induced sodium tail currents recorded from channels containing the L1014H mutation decayed 57-fold more rapidly than those carried by wild-type para/tipE channels. The effects of the F1538I mutation, identified in pyrethroid-resistant populations of B. microplus, have been assessed only in rat Nav1.4 sodium channels that were also mutated to enhance baseline pyrethroid sensitivity (Wang et al., 2001). Deltamethrin caused use-dependent modification of these rat Nav1.4 sodium channels. However, introduction of the F1538I mutation into these channels caused a loss of use-dependent modification by deltamethrin as well as a reduction in sensitivity to this compound. 5.1.4.5.3. Functional analysis of putative secondary mutations The effects of the T929I mutation, a putative second-site resistance mutation in P. xylostella and P. capitis, has been examined in para/tipE sodium channels expressed in oocytes (Vais et al., 2001). As a single mutation, the T929I substitution reduced the sensitivity of para/tipE channels to deltamethrin approximately tenfold. Deltamethrininduced tail currents carried by channels containing the T929I mutation decayed rapidly with first-order kinetics. Channels containing both the T929I and L1014F mutations, the combination found in highly resistant populations of Plutella xylostella were more than 10 000-fold resistant to deltamethrin when compared to wild-type channels. The effects of the E435K and C785R mutations, putative second-site mutations identified in several highly resistant B. germanica populations, on pyrethroid sensitivity have been examined in paraCSMA/tipE channels singly, together, and in combination with two primary resistance mutations. Channels containing the C785R mutation alone were identical to wild-type paraCSMA/tipE channels in their sensitivity to deltamethrin, but channels containing E435K single mutation and the E435K/ C785R double mutation were more sensitive to deltamethrin than wild-type channels (Tan et al., 2002b). When either the E435K or C785R mutations were combined with the L1014F resistance mutation, the resulting double mutants were approximately 20-fold less sensitive to deltamethrin than channels containing the single L1014F mutation and approximately 100-fold less sensitive than wild-type paraCSMA/tipE channels. Channels

p0305

p0310

p0315

Sodium Channels 17

containing all three resistance mutations, a situation found in highly resistant B. germanica populations, were more than 500-fold less sensitive to deltamethrin than wild-type channels. In a companion study, the E435K and C785R mutations were also evaluated as modifiers of deltamethrin resistance conferred by the V410M mutation (Liu et al., 2002). Insertion of either the E435K or C785R mutation into paraCSMA/tipE channels containing the V410M mutation did not significantly affect the level of resistance conferred by the V410M mutation, but channels containing all three resistance mutations were approximately sixfold less sensitive to deltamethrin than channels containing only the V410M mutation and 100-fold less sensitive than wild-type paraCSMA/tipE channels.

5.1.5. Conclusions 5.1.5.1. Unique Features of Insect Sodium Channels

Although sodium channel function and structure is strongly conserved between evolutionarily divergent animal species, the molecular characterization of insect sodium channels has identified several distinctive features. First, in contrast to the family of sodium channel a subunit genes in mammalian species, insects appear to have only a single gene (para in D. melanogaster and its orthologs) that encodes voltage-gated, sodium-selective channels that are involved in action potential generation. However, insects are capable of generating a remarkable diversity of sodium channel variants from this locus by alternative exon usage and RNA editing. The number of sodium channel variants generated by these posttranscriptional modifications found in any given species appears to be only a fraction of the theoretical maximum, and the patterns of exon usage do not appear to be strongly conserved between species. So far there is scant information on the functional significance of alternative exon usage. Insects also lack the sodium channel b subunit family of vertebrates and instead appear to use a novel family of glycoproteins, encoded by the tipE gene of D. melanogaster and its orthologs, as sodium channel auxiliary subunits. Genetic and cytochemical localization studies with D. melanogaster imply that the tipE protein is restricted in expression to the adult nervous system, suggesting that it may function as a modulator of sodium channel function and expression rather than as an obligatory auxiliary subunit. So far only two examples of tipElike genes have been described, but their broader existence in insects is inferred by the heterologous

coassembly of the D. melanogaster tipE protein with the para-orthologous sodium channel gene of B. germanica. It will be of interest to determine the breadth of distribution of tipE-like genes in the Insecta and in other animal taxa. Finally, insect genomes also contain a second sodium channel a subunit-like gene, dsc1 of D. melanogaster and its orthologs, that is equally divergent from the para-like sodium channel genes of insects and the vertebrate sodium channel a subunit gene family. The dsc1 protein is clearly not the sodium channel principally responsible for nerve action potential generation or the primary target for sodium channel-directed insecticides, and it may lack the stringent selectivity for sodium ions that is a hallmark of typical voltage-sensitive sodium channels. The functional significance of dsc1 and homologous proteins has remained obscure, but recent studies point to an as-yet undefined role in chemosensory pathways. 5.1.5.2. Sodium Channels and Pyrethroid Resistance

Molecular and genetic studies over the past decade have provided convincing evidence that point mutations in insect voltage-sensitive sodium channel genes are the primary cause of knockdown resistance to pyrethroids. Genetic linkage experiments and targeted DNA sequencing studies have consistently identified resistance-associated mutations in sodium channel genes that are orthologous to the para gene of D. melanogaster. Heterologous expression assays have documented the ability of many of these mutations, introduced into wild-type insect sodium channels either alone or in the combinations found in highly resistant populations, to reduce the sensitivity of expressed channels to pyrethroids exemplifying the type I and type II structural classes. Finally, the magnitude of resistance conferred by these mutations in heterologous expression assays in vitro has consistently been found to be in good agreement with the magnitude of resistance observed in resistant insect populations carrying the same mutations. These results not only identify the molecular mechanism of the kdr trait but also provide important confirmation that sodium channels encoded by the Vssc1 gene are the principal target site for the toxic actions of DDT analogs and pyrethroids. Although the actions of DDT and pyrethroids on sodium channels have been characterized in great depth and detail over the past three decades, numerous other sites of action for some or all of these insecticides have also been proposed (Soderlund and Bloomquist, 1989; Bloomquist, 1993). The toxicological impact of the knockdown

18 Sodium Channels

resistance mutations in para-orthologous sodium channels implies that actions of DDT and pyrethroids at this target alone are sufficient to account for the toxic effects of these compounds in whole insects. The search for sodium channel gene mutations in knockdown-resistant strains of various arthropod species has revealed the existence of resistance-associated polymorphisms at an unanticipated diversity of sites. Although not all of these polymorphisms have been shown to cause pyrethroid resistance, the multiplicity of sodium channel sequence polymorphisms associated with knockdown resistance contrasts with the situation for cyclodiene resistance in insects, which involves a mutation at a single amino acid residue in the subunit of the g-aminobutyric acid receptor–chloride ionophore encoded by the Rdl gene in all cases that have been examined (ffrench-Constant, 1994). The diversity of sequence polymorphisms that are potentially involved in knockdown resistance poses a significant challenge for the use of this information in pyrethroid resistance monitoring and management. The surprisingly large number of amino acid residues implicated as sites of resistance-causing mutations also poses challenges for the use of these mutations to map the pyrethroid binding site on the sodium channel. Not all of these residues are likely to interact physically with the pyrethroid molecule; mutations at some sites may indirectly alter the architecture of the pyrethroid site by altering or restricting the conformational flexibility of the sodium channel protein. As shown in Figure 3, most putative resistance mutations are located in the S5 and S6 helices and closely associated regions. Current models of sodium channel structure (Guy and Seetharamulu, 1986; Yellen, 1998; Lipkind and Fozzard, 2000) place the S5 and S6 domains in close proximity to each other adjacent to the inner pore of the channel. In this context, the clustering of a large number of resistance-associated mutations in the S5–S6 region of homology domain II implies an important role for this region of the sodium channel in determining, either directly or indirectly, the binding of pyrethroids. Recently, the crystal structure of a simple bacterial potassium channel was employed to generate a three-dimensional model of the S5–pore–S6 regions of the rat Nav1.4 voltage-sensitive sodium channel (Lipkind and Fozzard, 2000). The use of such a model to identify the spatial relationships among the resistanceassociated mutations in insect sodium channels may clarify the functional role of these residues in pyrethroid binding and action.

5.1.5.3. Future Exploitation of the Sodium Channel as an Insecticide Target

Despite the long and widespread use of DDT and then pyrethroids in the control of pests and disease vectors and the existence of knockdown resistance traits in populations of many pest and vector species, the pyrethroids remain an important and effective class of insecticides. Moreover, three key factors contribute to the continued value of the sodium channel as a target for future insecticides. First, toxicologically relevant sodium channels in insects are the products of a single gene and therefore exhibit conserved pharmacology in all neuronal tissues and insect life stages. Second, the value of sodium channel disruption as a mode of insecticidal action is amply demonstrated by the efficacy of sodium channel-directed natural toxins and insecticides. Third, toxin binding sites on the sodium channel other than the pyrethroid site (see Table 1) appear to be unaffected by mutations that confer pyrethroid resistance. In this context, each sodium channel binding domain can be envisioned as a separate potential target for insecticide discovery and development. These factors in favor of the continued exploitation of sodium channels for insect control are counterbalanced by the evolutionary conservation of sodium channel structure, function, and pharmacology across animal taxa. Intrinsic selectivity of agents for insect sodium channels is uncommon, and the development of novel insecticides directed toward this target is complicated by the potential for toxicity to nontarget species. The example of pyrethroids is instructive in this regard: the notable safety of pyrethroids for humans is based principally on differential metabolism rather than differential target sensitivity, and the use of pyrethroids has been limited in some contexts by undesirable toxicity to aquatic vertebrate and invertebrate species. The development of indoxacarb, which exploits differential metabolic bioactivation as a mechanism of selective toxicity, illustrates the potential for the discovery of novel sodium channel-directed insecticides that exhibit acceptable safety and selectivity.

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5.2 The Insecticidal Macrocyclic Lactones D Rugg, Fort Dodge Animal Health, Princeton, NJ, USA S D Buckingham and D B Sattelle, University of Oxford, Oxford, UK R K Jansson, Centocor, Malvern, PA, USA ß 2005, Elsevier BV. All Rights Reserved.

5.2.1. Discovery 5.2.2. Chemistry 5.2.2.1. Chemical Structure 5.2.2.2. Structure–Activity Relationships 5.2.3. Mode of Action 5.2.3.1. Biochemical and Molecular Action 5.2.3.2. Physiological Activity 5.2.3.3. Sublethal Toxicity 5.2.3.4. Metabolism 5.2.4. Biological Activity 5.2.4.1. Spectrum and Potency 5.2.4.2. Bioavailability 5.2.4.3. Safety and Selectivity 5.2.5. Uses 5.2.5.1. Crop Protection 5.2.5.2. Animal and Human Health 5.2.6. Resistance 5.2.6.1. Insects 5.2.6.2. Acarids 5.2.6.3. Helminths 5.2.7. Summary

5.2.1. Discovery The naturally occurring avermectins and milbemycins are fermentation products of actinomycetes in the genus Streptomyces. They are 16-membered, macrocyclic lactones, which have structural similarities to antibacterial macrolides and antifungal polyenes but lack their antifungal or antibacterial activities and do not inhibit protein or chitin synthesis (Fisher and Mrozik, 1989). Avermectins are produced by the soil microorganism, Streptomyces avermitilis MA-4680 (NRRL 8165), which was first isolated at the Merck Research Laboratories in 1976 from a soil sample collected near a golf course in Japan by researchers at the Kitasato Institute (Campbell et al., 1984). Milbemycins were first described from a culture of S. hygroscopicus and are structurally similar to the avermectins but lack the disaccharide substituent at C13 (Takiguchi et al., 1980). Mishima et al. (1975) first reported the acaricidal activity of milbemycins. The anthelmintic activity of these milbemycins was later elucidated following testing of individual members of the

25 26 26 27 27 27 30 31 31 32 32 35 36 38 38 39 39 39 41 42 42

milbemycin family (Mrozik et al., 1983). Milbemycins were also isolated from cultures of S. cyanogriseus in 1984 by workers at American Cyanamid and from S. thermoarchaensis by Glaxo researchers (Rock et al., 2002). Four homologous pairs of closely related compounds are produced through the fermentation of S. avermitilis MA-4680 (NRRL 8165), each pair comprising a major and minor component, usually produced in the approximate ratio of 8 : 2 (Lasota and Dybas, 1991). A mixture of one of these pairs, avermectin B1 (>80% B1a and 80%) and B1b (25 mg ml1) (Dybas, 1989; Lasota and Dybas, 1991; Jansson and Dybas, 1998). Although abamectin is toxic to certain aphids (e.g., LC90 values against black bean aphid, Aphis fabae and cotton aphid, Aphis gossypii range from 0.4 to 1.5 mg ml1) (Putter et al., 1981; Dybas and Green, 1984), it was not as effective at controlling aphids in translaminar assays (Wright et al., 1985). The reduced efficacy at controlling aphids is probably due to its prime activity as a stomach poison and its reduced concentrations in phloem tissue, where aphids actively feed. Poor residual control of aphids

in the field has been confirmed in several studies (Dybas, 1989 and references therein). Emamectin benzoate is a recently introduced avermectin that is highly potent to a broad spectrum of lepidopterous pests (Jansson and Dybas, 1998). LC90 values for emamectin benzoate against a variety of lepidopterous pests range between 0.002 and 0.89 mg ml1 (Dybas, 1989; Cox et al., 1995b; Jansson and Dybas, 1998). The hydrochloride salt of emamectin was up to 1500-fold more potent against armyworm species, e.g., beet armyworm (Spodoptera exigua), than abamectin (Trumble et al., 1987; Dybas et al., 1989; Mrozik et al., 1989). Emamectin hydrochloride was also 1720-, 884-, and 268-fold more potent to S. eridania than methomyl, thiodicarb, and fenvalerate, respectively, and 105- and 43-fold more toxic to cotton bollworm (H. zea) and tobacco budworm (H. virescens) larvae than abamectin (Dybas and Babu, 1988). Recent studies showed that emamectin benzoate was 875- to 2975-fold and 250- to 1300-fold more potent than tebufenozide to H. virescens and S. exigua, respectively. Emamectin benzoate was also 12.5- to 20-fold and 250- to 500-fold more potent than l-cyhalothrin and 175- to 400-fold and 2033- to 8600-fold more potent than fenvalerate to these two Lepidoptera, respectively (Jansson et al., 1998). Emamectin benzoate was 1.2- to 4.8orders of magnitude more potent to lepidopterous pests of cole crops (e.g., S. exigua, P. xylostella, and cabbage looper, Trichoplusia ni), than other new insecticides, including chlorfenapyr, fipronil, and tebufenozide (Jansson and Dybas, 1998; Argentine et al., 2002b). Emamectin benzoate is markedly less toxic to most nonlepidopteran arthropods. Emamectin is 8- to 15-fold less toxic to the serpentine leafminer, L. trifolii and the two-spotted spider mite, T. urticae, respectively, than abamectin (Dybas et al., 1989; Cox et al., 1995a). Emamectin benzoate and abamectin are comparable in their potency against Mexican bean beetle, Epilachna varivestis, and Colorado potato beetle, L. decemlineata (Dybas, 1989). Emamectin benzoate is markedly less toxic to black bean aphid, A. fabae, than abamectin (Jansson and Dybas, 1998). The macrocyclic lactones are widely used in animal health applications. The use of these compounds for antiparasitic therapy has been recently reviewed (Raymond and Sattelle, 2002; Vercruysse and Rew, 2002a). James et al. (1980) demonstrated the initial insecticidal activity of crude extracts and partially purified avermectins against Australian sheep blowfly, L. cuprina. Hughes and Levot (1990) examined the potencies of abamectin, ivermectin, and 4deoxy-4-epi(methyl amino) avermectin B1 in in vitro

The Insecticidal Macrocyclic Lactones

assays with L. cuprina. All three avermectins were equally potent against newly hatched larvae, and an order of magnitude more potent than diazinon when it was first introduced for blowfly control. Similarly, ivermectin is about two orders of magnitude more potent than deltamethrin against pyrethroidsusceptible larvae of L. cuprina (Rugg et al., 1995a). Ostlind et al. (1997) determined that ivermectin and emamectin were more potent than a number of common insecticides in a bioassay using larvae of Lucilia sericata. However, these workers found that abamectin was about 100-fold less toxic than ivermectin. Avermectins have similar potency against most myiasis producing dipterans. Abamectin or ivermectin concentrations of 5–20 ng ml1 in rearing medium were toxic to early stage larvae of Cochliomia macellaria (Chamberlain, 1982) and Chrysomya bezziana (Spradbery et al., 1985). Similarly, avermectins are highly potent against parasitic larvae in the families Cuterebridae (Ostlind et al., 1979; Roncalli, 1984), Gasterophilidae (Klei and Torbert, 1980; Craig and Kunde, 1981) and Oestridae (Preston, 1982; Yazwinski et al., 1983). Avermectins are also potent against hematophagous dipterans. Miller et al. (1986) demonstrated that the therapeutic anthelminthic (subcutaneous) dose of ivermectin would be effective against the blood-feeding horn fly, Haematobia irritans, for more than 2 weeks. Standfast et al. (1984) found that this same dose in cattle produced 99% mortality in engorged ceratopogonid midges 48 h after feeding for up to 10 days after treatment. In vitro studies have demonstrated that the subcutaneous dose of ivermectin in cattle kills 98% of buffalo flies fed for 7 days on blood taken 1 day after treatment (Kerlin and East, 1992). In vitro blood-feeding assays demonstrate that ivermectin is highly toxic to horn fly (88 h LC50 was 3–7 ng ml1) (Miller et al., 1986) and buffalo fly (H. irritans exigua) (48 h LC50 was 30–60 ng ml1) (Allingham et al., 1994). Adult tsetse fly (Langley and Roe, 1984), stable fly (Stomoxys calcitrans) (Miller et al., 1986), and mosquitoes (Pampiglione et al., 1985) tend to be less susceptible and generally survive feeding on animals treated at therapeutic dose rates. Fleas are generally considered to be relatively insensitive to avermectins or milbemycins systemically administered at, or slightly above, doses that provide ectoparasite control in production animals (Strong and Brown, 1987; Zakson-Aiken et al., 2001). However, Zakson-Aiken et al. (2001) examined a milbemycin and 19 avermectin aglycones, monosaccharides, and disaccharides for toxicity to fleas in an artificial membrane feeding system and

33

found that 19 of these compounds produced between 52% and 97% mortality in fleas fed a dose of 20 mg ml1. The potencies of these compounds, which included abamectin, selamectin, ivermectin, and milbemycin D, were remarkably similar, yet the most potent compound was inactive when tested at 1 mg kg1 subcutaneously in the dog (Zakson-Aiken et al., 2001). By contrast, selamectin provides effective flea control for up to 4 weeks after topical application at 6 mg kg1 on dogs (Banks et al., 2000; Bishop et al., 2000). Sucking lice on cattle are effectively controlled by therapeutic doses of avermectins, but biting lice are rarely affected by oral or injectable doses (Benz et al., 1989). However, topical administration to cattle does give effective control of biting lice (Benz et al., 1989). In vitro studies with sheep biting lice indicate that efficacy is not due to direct contact activity, but rather to the differential distribution following application resulting in the availability of a toxic oral dose. Rugg and Thompson (1993) examined the toxicity of abamectin, ivermectin, and cypermethrin to the sheep-biting, louse Damalinia ovis. These workers found that avermectins were an order of magnitude less potent than cypermethrin when assessed in a contact bioassay. However, in an assay incorporating ingestion and contact action both macrocyclic lactones were an order of magnitude more potent than cypermethrin. Macrocyclic lactones are also highly toxic to many dung-feeding insects, but toxicity may vary widely between life stages. Dung-feeding Diptera and Coleoptera are highly susceptible; however, adult dung beetles are relatively insensitive. Rugg (1995) found that adult Onthophagus gazella were four orders of magnitude less susceptible to ivermectin or moxidectin than larvae in in vitro bioassays. This relative insensitivity in adult dung beetles with no significant effects on adult mortality seen in mature beetles fed dung from cattle treated with ivermectin has been reported previously (RidsdillSmith, 1988; Wardhaugh and Rodriguez-Mendez, 1988; Fincher, 1992). However, newly emerged and immature adult beetles may be affected negatively by ivermectin residues (Wardhaugh and Rodriguez-Mendez, 1988; Houlding et al., 1991). Adult dung beetles are sexually immature at emergence and have an initial period of intense feeding. It is during this period that they would be most susceptible to residues in feces. Nevertheless, the relative insensitivity of adult dung beetles is attributable to ingestion being the main route of toxicity, to the feeding behavior of the beetles, and the characteristics of the compounds. Adult dung beetles ingest only the liquid and colloidal constituents of dung,

34 The Insecticidal Macrocyclic Lactones

as their mouthparts are unsuitable for solids (Bornemissza, 1970). Avermectins and similar compounds bind tightly to soil and organics, and are virtually insoluble in water (Halley et al., 1989b) and could be expected to partition differentially to the solid components of dung. Thus, adults would be expected to ingest relatively low concentrations of these compounds when feeding on the liquid portion of dung. Milbemycins generally tend to have lower toxicities to insects than the avermectins, but similar or higher potencies against some nematodes and acarids (McKellar and Benchaoui, 1996). In vitro bioassays have shown that moxidectin is 50- to 100-fold less toxic than ivermectin and abamectin to the sheep blowfly, L. cuprina, and about sixfold less potent than these two avermectins against sheepbiting lice (Rugg, 1995). This difference is clearly demonstrated in the effects of these compounds on dung feeding insects. An in vitro comparison of moxidectin and abamectin (Doherty et al., 1994) found that abamectin was about 50-fold more toxic to larvae of a dung beetle (O. gazella) and buffalo fly than moxidectin when added to cattle feces. Rugg (1995) observed a similar result with both ivermectin and abamectin, which were at least an order of magnitude more potent than moxidectin to adults and larvae of a dung beetle and larvae of a dung-feeding fly (T. brevicornis). While ivermectin residues are generally considered to be toxic to some dung beetle larvae and fly larvae for up to 4 weeks after the treatment of cattle, there are differing reports for moxidectin. Strong and Wall (1994) found that moxidectin residues had no toxic effects on the larvae of beetles and cyclorrhaphous flies in cattle dung, and similarly, Fincher and Wang (1992) concluded that dung from moxidectin treated animals did not affect two species of dung beetles. However, Webb et al. (1991) reported that feces from cattle treated with a low, controlledrelease dose of moxidectin were toxic to larvae of the face fly, Musca autumnalis, and concluded that the therapeutic injectable dose of moxidectin would be an effective control of M. autumnalis larvae in dung. Miller et al. (1994) reported significant mortality in horn fly larvae in dung for up to 28 days after oral or injectable treatment of cattle with moxidectin. Rank ordering of the negative effects of various avermectins and moxidectin on insect development in the dung of treated cattle was doramectin >ivermectin  eprinomectin  moxidectin (Floate et al., 2001, 2002). While moxidectin is generally less potent against insects than the avermectins in in vitro evaluations, it is commonly used at the same or similar therapeutic

dose rates in production animals. Interestingly, the spectrum of insects controlled by these products is remarkably similar (Shoop et al., 1995; McKellar and Benchaoui, 1996). The only macrocyclic lactones that have been specifically developed for primary insect control on animals are avermectins: ivermectin for blowfly and lice control on sheep (Eagleson et al., 1993a, 1993b; Rugg et al., 1993, 1995a, 1995b; Thompson et al., 1994), and selamectin for flea control on cats and dogs (Benchaoui et al., 2000; McTier et al., 2000a, 2000b). 5.2.4.1.2. Acarids, copepods, and helminths Macrocyclic lactones are highly potent acaricides. Abamectin and natural milbemycins are widely used as agricultural miticides. Abamectin is highly effective against a broad spectrum of phytophagous mites and is one of the most potent acaricides with LC90 values ranging between 0.02 and 0.24 mg ml1 for mites in the families Tetranychidae, Eriophyidae, and Tarsonemidae (Lasota and Dybas, 1991). Eriophyid mites, such as the citrus rust mite, Phyllocoptruta oleivora, and various tetranychid mites, such as T. urticae, are among the most susceptible (Dybas, 1989). Other phytophagous mites, such as citrus red mite, Panonychus citri (LC90 ¼ 0.24 mg ml1) are slightly more tolerant (Dybas, 1989). Similarly, ticks and parasitic mites are variably susceptible to the macrocyclic lactones. Much of the variation in potency against ticks relates to the different life cycle of ticks, the relatively slow action of these compounds against ticks, and the interactions with bioavailability of the compound in the host. These compounds tend to be much more effective against single host ticks; and are often considered relatively poor compounds against multihost ticks (McKellar and Benchaoui, 1996). In multihost ticks, adults are the infestive stage and are the least sensitive to macrocyclic lactones. Parasitic mites show similar variation: the deep burrowing sarcoptic mange mites that feed directly on blood and body fluids are highly sensitive, while chorioptic mange mites that live and feed on the superficial dermal surface are relatively insensitive (Benz et al., 1989). Macrocyclic lactones are also toxic to a number of parasitic copepods and ivermectin, emamectin benzoate, and doramectin have been used for control of parasitic sea lice in farmed salmon (Burka et al., 1997; Davies and Rodger, 2000; Roth, 2000). The macrocyclic lactones are highly potent nematicides, but have no useful activity against trematodes or cestodes. These compounds are especially toxic to the microfilarial stages of many filaroid nematodes (McKellar and Benchaoui, 1996). This extreme potency against microfilaria is best

The Insecticidal Macrocyclic Lactones

demonstrated in the use of these compounds for heartworm control in dogs. Oral doses of 3–12 mg kg1 of moxidectin or ivermectin once a month provide effective protection against heartworm infection in dogs (Guerrero et al., 2002). The oral use rate of these compounds for general nematode control in ruminants is about 200 mg kg1 (McKellar and Benchaoui, 1996). 5.2.4.2. Bioavailability

5.2.4.2.1. Crop protection uses Macrocyclic lactones are very susceptible to rapid photodegradation. The half-life of abamectin as a thin film under artificial light or simulated sunlight was 4–6 h and the rate of degradation of abamectin on petri dishes and on leaves was markedly greater in light than in dark environments (MacConnell et al., 1989). The half-life of emamectin benzoate has been estimated to be 0.66 days on celery (Feely et al., 1992) and is expected to be even shorter on cole crops. Photoirradiation of avermectins leads to the production a large number of decomposition products (Mrozik et al., 1988), and a number of these photodegradates have also been identified for abamectin (Crouch et al., 1991) and emamectin benzoate (Feely et al., 1992). Many of these photodegradates show impressive insecticidal activity against larval lepidopterans (Argentine et al., 2002a). Despite the rapid degradation of avermectin insecticides in sunlight, significant amounts of these compounds are taken up rapidly via translaminar movement into sprayed foliage (Wright et al., 1985; Dybas, 1989 and references therein). When abamectin was applied to plants held in the dark, the prolonged stability resulted in greater penetrability into leaves and more effective mite control (MacConnell et al., 1989). Wright et al. (1985) demonstrated that this translaminar movement of abamectin produced good control of mites, but aphids were not controlled. They surmised that abamectin residues were probably abundant in the parenchyma tissues of leaves where mites feed, whereas abamectin probably did not distribute into the phloem tissue where aphids feed. The excellent residual efficacy of abamectin against several dipteran and lepidopteran leafminers is probably also due to translaminar movement and the presence of abamectin reservoirs in parenchyma tissue (Jansson and Dybas, 1998). This lack of true systemic activity (distribution into the phloem) in plants following spray application is demonstrated by the lack of residual efficacy against aphids. Spray applications of abamectin that had good contact efficacy provided little if any residual control of the aphids Myzus persicae (Johnson, 1985) and A. fabae

35

(Green and Dybas, 1984). Similarly, the lack of downward transport of abamectin sprayed on foliage results in poor control of root knot nematodes (Stretton et al., 1987; Cayrol et al., 1993). By contrast, root dip, soil application or stem injections of avermectins have shown the potential to provide effective control of these parasites (Sasser et al., 1982; Jansson and Rabatin, 1997, 1998). 5.2.4.2.2. Animal health uses In animal health, the high potency and lipophilic nature of the macrocyclic lactones allows extensive systemic delivery through a variety of routes. These compounds are used in oral, injectable, and topical formulations. The route of delivery, however, influences the bioavailability and distribution of the compounds in the host animal and thus the efficacy against different target parasites. Similarly, bioavailability can vary dramatically between species and be markedly affected by feed quantity (Ali and Hennessy, 1996) or composition (Taylor et al., 1992) when orally administered. Efficacy against parasites may be dependent on transport in the plasma and distribution to vascular or nonvascular sites that constitute the parasite habitat. The pharmacokinetics of the macrocyclic lactones were reviewed by Hennessy and Alvinerie (2002). Following most means of administration, macrocyclic lactones are eliminated fairly rapidly from the treated animals; elimination half-lives in various species range from about 3 to 10 days (see Hennessy and Alvinerie, 2002). Regardless of the means of administration or the species treated, the main route of elimination of the compounds is via feces with the compounds passing into the gastrointestinal tract in the bile (Hennessy and Alvinerie, 2002). A number of controlled release devices (Shoop and Soll, 2002) or sustained release formulations (Rock et al., 2002; Shoop and Soll, 2002) using macrocyclic lactones have been used to provide long-term protection against parasites. The most dramatic differences between application methods are seen with certain insect ectoparasites. While a number of blood-sucking parasites, such as sucking lice of cattle, are controlled by therapeutic doses of macrocyclic lactones administered orally or by injection, chewing lice are generally not susceptible. Topical applications (generally at about a 2.5-fold higher dose rate) are usually highly effective against chewing lice (Strong and Brown, 1987; Vercruysse and Rew, 2002b). Similar results have also been observed in lice on sheep (Coop et al., 2002). While the efficacy and spectrum of these topical formulations against internal parasites is similar to the oral or injectable treatments, this

36 The Insecticidal Macrocyclic Lactones

greater efficacy against chewing lice is thought to be a result of a high concentration of the active ingredient on the surface of the skin where these parasites feed (Titchener et al., 1994). Similarly, most of the macrocyclic lactone pour-on formulations for cattle have demonstrated a persistent efficacy against the blood-feeding horn fly and buffalo fly that is not seen with oral or injectable formulations (Vercruysse and Rew, 2002b). This apparent greater efficacy of pour-on formulations against external parasites is presumably the result of deposition of the active compound on, and in, the skin. The high lipophilicity of the macrocyclic lactones is likely to result in the formation of depots of active ingredient in skin lipids and oil and fat secretions. This characteristic has been exploited in topical formulations of ivermectin that provide long-term residual control of blowfly and lice on sheep (Eagleson et al., 1993a, 1993b; Thompson et al., 1994), and selamectin which controls fleas on dogs and cats for at least 1 month (Benchaoui et al., 2000). Even greatly exaggerated oral or injectable doses of macrocyclic lactones are considered unlikely to provide residual control of fleas (Zakson-Aiken et al., 2001). In sheep, topically applied ivermectin is thought to bind to skin lipids and secretions and may be passively distributed around the sheep’s body in this medium following application to a discrete site (Rugg and Thompson, 1997). Selamectin is thought to selectively partition into sebaceous glands of dogs and cats from where it is slowly released to the skin over an extended period (Hennessy and Alvinerie, 2002). 5.2.4.3. Safety and Selectivity

5.2.4.3.1. Crop protection uses Abamectin is generally considered to be nonphytotoxic under normal use even in sensitive ornamental plants (Wislocki et al., 1989), although mild foliar spotting has been noted in some ferns, daisies, and carnations (Dybas, 1989). Phytotoxicity was not affected by the addition of spray oils (Green et al., 1985), although, the addition of spray oils to abamectin has produced epidermal damage in pears that is not seen with the use of abamectin alone (Hilton et al., 1992). Integrated pest management (IPM) involves the use of all available means (chemical, biological, cultural, physical, etc.) to achieve effective control of pests or pest damage. Central to the design of IPM programs are selective insecticides that reduce pest abundance and yet are innocuous to beneficial predators or parasites by allowing their survival and range expansion. The suitability of pesticides for use in IPM programs is based on the differential toxicity of a compound to pest and beneficial arthropod

populations. Differential toxicity can occur through pharmacokinetic or metabolic differences between pests and beneficial organisms (physiological selectivity) or by a compound’s unique qualities that result in toxic exposure of phytophagous pests with concomitant reduced exposure of beneficial organisms (ecological selectivity). The macrocyclic lactone insecticides possess both of these qualities, thereby making them highly selective and compatible with IPM. The ecological selectivity of the avermectins and milbemycins results from a number of qualities of these compounds following application in the field. As noted earlier, avermectin insecticides are readily taken up by plant foliage. Foliar uptake via translaminar movement results in a reservoir of the toxicant inside the leaf (MacConnell et al., 1989). Any remaining compound on the outside of foliage is rapidly degraded by sunlight, thereby resulting in minimal residues of the compound on the plant surface soon after application. These qualities of the avermectin insecticides that result in the selective availability of the compounds to phytophagous pests and reduced exposure to nonphytophagous organisms are the main reasons for their compatibility with beneficial arthropods and IPM programs. A number of researchers have shown that applications of abamectin did not disrupt the Liriomyza leafminer–parasitoid complex on a variety of vegetable crops (see Dybas, 1989). Trumble and Alvarado Rodriguez (1993) demonstrated that because of the reduced toxicity of abamectin to natural enemies, it was an important component of a multiple-pest IPM program on fresh market tomato that was based on intensive sampling, parasitoid releases, mating disruption, microbial pesticides, and abamectin. Abamectin has also been shown to be less detrimental to certain life stages of two leafminer parasitoids, Diglyphus intermedius and Neochrysocharis punctiventris, compared with other commonly used insecticides (Schuster, 1994). The differential toxicity of abamectin to a variety of pest and predatory mite species is also well documented (see Lasota and Dybas, 1991). Hoy and Cave (1985) showed that although abamectin was toxic to the predator Metaseiulus occidentalis, it was more toxic to the two-spotted spider mite, T. urticae. Similar results have been found with other predatory mite species (Grafton-Cardwell and Hoy, 1983; Zhang and Sanderson, 1990). Hoy and Cave (1985) also showed that 2–4 day fieldaged residues of abamectin were safe for M. occidentalis. Several other studies demonstrated that abamectin was less toxic to naturally occurring

The Insecticidal Macrocyclic Lactones

and introduced natural enemies of pest mites, and for this reason, it selectively kills target pests and conserves their natural enemies (Dybas, 1989; Lasota and Dybas, 1991). Zchorifein et al. (1994) found that abamectin was compatible with Encarsia formosa, an important component of IPM programs for greenhouse whitefly, Trialeurodes vaporariorum. No mortality was observed when adult parasitoids were exposed to 24 h residues of abamectin. They found that a management program based on the combined treatment of abamectin with releases of E. formosa maintained lower densities of the whitefly on poinsettia throughout the season and required fewer applications of abamectin compared with a management program based on applications of abamectin alone. Similarly, Brunner et al. (2001) found that while abamectin was highly toxic when applied topically to two parasitoids of leafrollers, 1-day-old residues were not toxic. Like abamectin, emamectin benzoate is less toxic to beneficial arthropods such as honeybees, parasitoids, and predators. Kok et al. (1996) reported that emamectin hydrochloride had minimal adverse effects on two hymenopterous parasitoids (Pteromalus puparum and Cotesia orobenae) of Lepidoptera on broccoli. Contact activity of emamectin benzoate residues on alfalfa against two species of bees declined rapidly and was negligible 24 h after treatment (Chukwudebe et al., 1997). They determined that bee mortality was directly related to transient dislodgeable residues and thus these beneficial insects could be expected to survive and colonize treated crops within relatively short intervals after applications of emamectin benzoate. Sechser et al. (2003) found that emamectin benzoate was relatively safe to a number of insect predators of sucking pests on cotton. The selectivity of abamectin has been widely investigated in mites. Trumble and Morse (1993) showed that abamectin was compatible with Phytoseiulus persimilis in field-grown strawberry. They found that the highest net economic return was achieved when several releases of the predatory mite P. persimilis were integrated with suitably timed applications of abamectin. In a laboratory leaf-disc bioassay, adult females of the predatory mite Amblyseius womersleyi dipped in abamectin solution showed low mortality (16.6%), while all T. urticae females died within 24 h after dipping (Park et al., 1996). In apple, abamectin was highly compatible with the phytoseiid mites M. occidentalis and Amblyseius fallacis. Abamectin was effective at reducing and maintaining numbers of European red mites (Panonychus ulmi) at or below the

37

economic threshold for at least 3 weeks after application. Although predatory mite numbers declined initially, they quickly resurged resulting in favorable predator to prey ratios that were comparable or superior to those found in nontreated plots, and aided in mite suppression for the remainder of the growing season (Jansson and Dybas, 1998). In glasshouse roses, Sanderson and Zhang (1995) found that full canopy applications of abamectin at 3–5 day spray intervals effectively controlled T. urticae populations, but were detrimental to the predatory mite P. persimilis. However, by applying abamectin to the upper canopy of plants only, excellent mite control in the marketable portion of the plant was achieved and predatory mite populations in the lower canopy were conserved. This demonstrated the potential for integrating abamectin with natural predators for control of a target pest, even under intense abamectin pressure. 5.2.4.3.2. Animal health uses The macrocyclic lactones are generally considered to be relatively safe in mammals. Glutamate-gated chloride channels have not been reported in mammals. Clinical signs are associated with neurotoxicity of the central nervous system as this is the site of GABA-gated chloride channels in mammals. The poor distribution of these compounds through the blood–brain barrier and/or their rapid removal by the transmembrane protein, P-glycoprotein, are thought be responsible for their lack of toxicity to mammals (Shoop and Soll, 2002; Abu-Qare et al., 2003). Increased susceptibility to avermectins has been observed in Murray Grey cattle (Seaman et al., 1987) and in some collie-breed dogs (Paul et al., 1987). It is thought that a deficiency of P-glycoprotein in these animals results in increased accumulation of avermectins in the central nervous system and the resultant neurotoxicity (Shoop and Soll, 2002; Roulet et al., 2003). In animal health, nontarget effects of macrocyclic lactones are generally minimized due to their main use as systemic parasiticides. The compounds are applied discretely to the target host and not released into the environment. The major nontarget effects in animal health uses are on the insect fauna of dung of treated animals. Regardless of the route of administration, the major means of elimination of these compounds from animals is via the feces and the major component in feces is usually the parent compound (Campbell et al., 1983; Chiu and Lu, 1989). Feed-through compounds for the control of pestiferous Diptera in dung have been in use, especially in cattle, for many years, yet little attention has been paid to the effects on nontarget dung

38 The Insecticidal Macrocyclic Lactones

fauna (Strong, 1992). Similarly, most of the compounds used as external parasiticides for cattle are toxic to dung insects, but organophosphorus and pyrethroid compounds are more toxic to dunginhabiting insects than avermectins (Bianchin et al., 1992). However, there has been a great deal of attention given to possible environmental consequences of systemic parasiticides, especially avermectins and related compounds, with entire journal volumes (e.g., Veterinary Parasitology 48, 1993) and dedicated regulatory or scientific workshops (National Registration Authority, 1998; Alexander and Wardhaugh, 2001) devoted to this subject. This rise in interest is probably related to the widespread use of these compounds. Their broad spectrum, encompassing both nematodes and arthropods, has resulted in their extensive use, and especially where gastrointestinal worms are the prime target, most arthropods could be considered nontarget organisms. Certainly, in countries where the introduction of dung beetles to control dung build-up and dung breeding pests have occurred, any compounds with adverse effects on these beneficial agents will gain some attention. Also, the differing toxicities of related compounds marketed by different companies may be perceived as providing a commercial advantage. Hence, authors have reported the effects of ivermectin in controlling dung-breeding pestiferous Diptera (e.g., Meyer et al., 1980; Miller et al., 1981; Marley et al., 1993), or its toxicity to dung beetles (e.g., Ridsdill-Smith, 1988; Sommer and Overgaard Nielsen, 1992; Sommer et al., 1992, 1993), and that dung degradation may be retarded (e.g., Wall and Strong, 1987; Sommer et al., 1992) or not affected (e.g., Jacobs et al., 1988; McKeand et al., 1988; Wratten et al., 1993). Other reports have appeared directly comparing the effects of avermectins and moxidectin on dung fauna (Fincher and Wang, 1992; Doherty et al., 1994; Floate et al., 2001, 2002). There is little doubt that avermectins and related compounds have the potential to affect dunginhabiting insects. Moxidectin has little if any impact on dung insects, but also is less efficacious against pestiferous fly larvae. The selection of these compounds for certain animal health uses should be evaluated on the basis of their efficacy against pest target and on concomitant environmental concerns. Thus, for example, ivermectin use in a controlledrelease device for sheep provides added protection against breech strike (Rugg et al., 1998a), and residues in dung likely reduce populations of the adult blowfly population (Cook, 1991, 1993), while exerting little impact on the rate of degradation of sheep dung where insects play a relatively minor role

(Cook, 1993; King, 1993). Where there are concerns over possible adverse effects of treatment for gastrointestinal worm infestation on cattle dung fauna, moxidectin would be the compound of choice (Strong and Wall, 1994). If control of arthropod ectoparasites or their dung-feeding stages is of importance (Marley et al., 1993), an avermectin may be preferred because of its greater toxicity to insects. Even so, a study examining the effects of commercial ivermectin use in cattle in South Africa (Scholtz and Kru¨ ger, 1995) found that dung insect populations recovered rapidly following an initial depression. They concluded that the long-term effects on dung fauna might not be as severe as previous studies have claimed. However, simulation of the impact of eprinomectin on dung beetle populations using computer modeling suggests that, in the absence of immigration, a single treatment of eprinomectin could reduce beetle activity by 25–35% in the next generation (Wardhaugh et al., 2001). In studies directly comparing the effects of macrocyclic lactones in cattle dung, avermectins result in lethal and sublethal effects on dung fauna in feces excreted for a number of weeks after treatment, while moxidectin residues are relatively innocuous (Fincher and Wang, 1992; Doherty et al., 1994; Floate et al., 2001, 2002; Wardhaugh et al., 2001).

5.2.5. Uses 5.2.5.1. Crop Protection

Abamectin is registered worldwide for control of mites and certain insect pests on a variety of ornamental and horticultural crops and on cotton. Abamectin is effective for controlling agromyzid leafminers, Liriomyza spp., Colorado potato beetle, L. decemlineata, diamond back moth, P. xylostella, tomato pinworm, K. lycopersicella, citrus leafminer, Phyllocnistis citrella, and pear psylla, Cacopsylla pyricola. Abamectin has also been incorporated into bait to control red imported fire ants and cockroaches. These and other urban pest applications were reviewed by Lasota and Dybas (1991). Recent studies have shown that avermectins have potential as a component in Africanized honeybee abatement programs (Danka et al., 1994). Another novel use is the injection of abamectin into trees which controls certain phytophagous arthropods, such as elm leaf beetle (Pyrrhalta luteola), for up to 83 days after application (Harrell and Pierce, 1994). The main commercial agricultural use of macrocyclic lactones is as acaricides. These compounds have demonstrated mite control and residual activity superior to most acaricides on a variety of ornamental crops, food crops, and cotton (Putter

The Insecticidal Macrocyclic Lactones

et al., 1981; Dybas and Green, 1984; Dybas, 1989). Macrocyclic lactones are also highly effective for the control of plant parasitic nematodes following injection application (Jansson and Rabbatin, 1997, 1998; Takai et al., 2003) or soil incorporation (Blackburn et al., 1996). Emamectin benzoate, milbemectin, and nemadectin are used commercially for the control of pine wilt nematode in Japan. Emamectin benzoate is used for broad-spectrum control of lepidopteran pests on certain vegetable crops. This compound is very effective against numerous lepidopteran pests of a variety of crops. It is expected that the compound will be used to control lepidopterous pests on a wide variety of vegetable, tree, and row crops and also for control of thrips on eggplant and tea in Japan (Jansson and Rabatin, 1997). 5.2.5.2. Animal and Human Health

The macrocyclic lactones have had a dramatic impact on animal health. Their potency and broad spectrum has resulted in this chemistry dominating the parasiticide market. Macrocyclic lactones have been commercialized for the control of nematodes and arthropod parasites for most common foodproducing and companion animals and are widely used ‘‘off-label’’ for parasite control in many other species. Avermectins (e.g., emamectin) are also used for control of copepod parasites of farmed salmon (Davies and Rodger, 2000). In human health, ivermectin has been used since 1987 in a compassionate program in Africa and Central and South America to control Onchocerca volvulus which is the causative agent of river blindness in man (Shoop and Soll, 2002). The program was expanded recently to include the reduction of spread of elephantiasis caused by lymphatic filarid nematodes. Recently, moxidectin has been demonstrated to be safe and well tolerated in humans (Cotreau et al., 2003). Moxidectin is more efficacious than ivermectin in Onchocerca models and is currently undergoing clinical trials prior to inclusion in filarid control programs (Molyneux et al., 2003). Ivermectin has been approved for treatment of intestinal strongiloidosis scabies in humans, and is also useful for the treatment of louse infestations (Elgart and Meinking, 2003). These authors consider that there may be a number of additional uses for the macrocyclic lactones as antiparasitic medications for parasites of skin in humans.

5.2.6. Resistance 5.2.6.1. Insects

Resistance to the avermectins in insects has been reviewed by Clark et al. (1995). In general, a variety

39

of biochemical and pharmacokinetic mechanisms may contribute to avermectin resistance in arthropods, although biochemical mechanisms tend to be more important among most arthropod systems studied to date. However, there are many conflicting reports on the importance of different resistance mechanisms. Similarly, cross resistance between avermectin insecticides/acaricides and other classes of chemistry is not well understood, and there are probably equal numbers of reports demonstrating cross-resistance as there are those refuting it. Cross-resistance between abamectin and pyrethroids has been reported for field strains of houseflies (Scott, 1989; Geden et al., 1990). Abro et al. (1988) proposed that there was a low level of cross resistance to abamectin in a field strain of diamondback moth that was highly resistant to DDT, malathion, and cypermethrin when assessed in a topical assay. However, the same population was equally susceptible to abamectin as the baseline colony in an ingestion bioassay. Roush and Wright (1986) found no evidence of cross-resistance to avermectins in houseflies resistant to diazinon, dieldrin, DDT, or permethrin, and Parella (1983) reported no crossresistance to avermectins in a pyrethroid-resistant agromyzid fly. Campanhola and Plapp (1989) determined that there was no cross-resistance to abamectin in pyrethroid-resistant tobacco budworm, where both target site resistance and metabolic resistance to pyrethroids were present. Similarly, Cochran (1990) concluded that pyrethroid-resistant field populations of German cockroach were not crossresistant to abamectin. Argentine and Clark (1990) examined a susceptible laboratory strain and a multiple-resistant field strain of Colorado potato beetle, and found no difference in their dose responses to abamectin. No cross-resistance to abamectin was observed in a laboratory-selected pyrethroid-resistant strain of citrus thrip, although cross-resistance was detected to DDT and some organophosphates and carbamates (Immaraju and Morse, 1990). Rugg and Thompson (1993) reported that pyrethroid-resistant sheep biting lice (B. ovis) were not cross-resistant to ivermectin. Beeman and Stuart (1990) examined a number of pesticides against a field-collected strain of red flour beetle (Tribolium castaneum) resistant to lindane that had been further selected with dieldrin in the laboratory. These workers found cross-resistance to other cyclodienes, but not to abamectin nor to organophosphates. Resistance to cyclodienes, accounted for by channel mutants in the Rdl GABA-gated chloride channel subunit (review: ffrench-Constant, 1994), usually extends to all insecticides that block chloride ion channels, and this resistance is not

40 The Insecticidal Macrocyclic Lactones

conferred to avermectins as these activate chloride channels and therefore act at a separate site (Bloomquist, 1993). Rohrer et al. (1995) showed that fipronil, which interacts with GABA-gated chloride channels (as an antagonist) did not affect ivermectin binding either in Drosophila head membranes or in locust neuronal membranes. Ismail and Wright (1991) examined the toxicities of a number of insect growth regulators, and abamectin, to a susceptible strain of diamondback moth and to strains selected for resistance to the growth regulators chlorfluazuron and teflubenzuron. They found that cross-resistance to other growth regulators was variable and did not detect cross-resistance to abamectin. Recently, Zhao et al. (2002) showed that the resistance to spinosad in a field population of diamondback moth was not cross-resistant to emamectin benzoate. Scott (1989) proposed that cross-resistance to abamectin in pyrethroid-resistant houseflies was polygenic and due to decreased cuticular penetration and enhanced metabolism mediated by mixedfunction oxidases. Subsequently, Scott et al. (1991) reported laboratory selection of high-level abamectin resistance in these field-collected houseflies; after seven selections resistance ratios of >60 000fold (determined by topical application) and 36-fold (determined by residual exposure) were reached. In their study the synergists piperonyl butoxide, diethyl maleate, and S,S,S-tributylphosphorotrithioate did not affect the toxicity of abamectin, indicating that resistance was not due to enhanced metabolism mediated by mixed function oxidases, glutathione transferases, or hydrolases. Konno and Scott (1991) found that resistance in this selected strain resulted from decreased cuticular penetration (>1700-fold) and altered abamectin binding (35-fold). This strain was also cross-resistant to two abamectin analogs and resistance was highly recessive. Laboratory selection of abamectin against two strains of Colorado potato beetle resulted in resistance levels of 21-fold for a mutagen-induced resistance and 38-fold by direct selection. Both resistance factors were incompletely dominant and polyfactorial (Argentine and Clark, 1990). Oxidative metabolism and possibly carboxylesterase activity were responsible, in part, for resistance to abamectin (Argentine, 1991; Argentine et al., 1992). Lamine (1994) found that abamectin-resistant larvae of L. decemlineata were seven- to tenfold less susceptible to emamectin benzoate than the baseline colony in a topical bioassay. No cross-resistance between abamectin and emamectin benzoate was evident in adult beetles. Recent studies showed

that L. decemlineata populations which were 15to 23-fold less sensitive to abamectin in a contact, topical bioassay (Clark et al., 1995) were classified as susceptible to abamectin in a diet-based ingestion assay (Dively and Jansson, 1996). Recently, oxidative metabolism was confirmed as a principal resistance mechanism in laboratory-selected, abamectin-resistant Colorado potato beetle (Yoon et al., 2002; Gouamene-Lamine et al., 2003). Hughes and Levot (1990) reported that a laboratory-selected strain of L. cuprina resistant to organophosphates and carbamates exhibited a low level cross-resistance (two- to threefold) to abamectin and ivermectin when compared to susceptible flies. However, emamectin benzoate was equally toxic to both strains. This strain also exhibited a low level of cross-resistance to pyrethroids (Sales et al., 1989). A laboratory-selected pyrethroid-resistant strain of L. cuprina that showed cross-resistance between an organophosphate and a carbamate insecticide (Sales et al., 1989) was subsequently selected with deltamethrin, diflubenzuron, and butacarb resulting in levels of resistance in excess of 1000-fold to all three compounds (Kotze and Sales, 1994). These workers also found that metabolic inhibitors significantly synergized the three selected strains, indicating the involvement of both monooxygenases (mixed-function oxidases) and esterases in these resistances. Rugg et al. (1995a) showed that the same pyrethroid- and carbamate-resistant laboratory strain of L. cuprina was not cross-resistant to ivermectin. Organophosphate-resistant field strains of L. cuprina have also been shown to have no crossresistance to avermectins (Hughes and Levot, 1990; Rugg et al., 1998b). Rugg et al. (1998b) selected larvae of L. cuprina with ivermectin for over 60 generations and achieved an eightfold increase in tolerance compared to the parental strain. This low-level resistance reverted rapidly in the absence of selection pressure, and although enzymatic metabolism was implicated, no specific resistance mechanisms were determined. Kotze (1995) found that the induction of monooxygenase and glutathione transferase enzymes with phenobarbital in insecticide-susceptible L. cuprina larvae resulted in increased tolerance to butacarb, diazinon, and diflubenzuron. In similar experiments, Rugg et al. (1998a) found no increase in tolerance to ivermectin in induced flies that had reduced susceptibility to diazinon. Resistant field populations of P. xylostella from Malaysia were not cross-resistant to Bacillus thuringiensis (Iqbal et al., 1996). Lasota et al. (1996) showed that there was no cross-resistance between abamectin, emamectin benzoate, permethrin, and

The Insecticidal Macrocyclic Lactones

methomyl in P. xylostella using an ingestion bioassay. Following laboratory selection of a field population of P. xylostella from China, testing of F1 progeny from reciprocal crosses between abamectinresistant and abamectin-susceptible strains indicated that resistance was autosomal and incompletely recessive and backcross studies suggested that the resistance was probably controlled by more than one gene (Liang et al., 2003). Further, there was little cross-resistance between abamectin and four pyrethroids (deltamethrin, b-cypermethrin, fenvalerate, and bifenthrin) and no cross-resistance between abamectin and the acylureas chlorfluazuron or flufenoxuron. Morse and Brawner (1986) reported LC50 values ranging from 7 to 21 mg l1 in strains of citrus thrips that had not been exposed to abamectin. Immaraju and Morse (1990) retested one of these strains that had been maintained in the laboratory and also selected for resistance to fluvalinate, and determined LC50 values of 0.9 and 0.4 mg ml1, respectively. Responses to abamectin in nonexposed populations varied by over 50-fold suggesting that thrips may have a large inherent variation in susceptibility to abamectin. Immaraju et al. (1992) reported the occurrence of two abamectin-resistant populations of western flower thrips in Californian glasshouses. Resistance ratios (LC90) of 18- and 798-fold were attributed to intensive use of abamectin during the previous year (five or ten applications, respectively). In summary, resistance to avermectins has been demonstrated in a number of insects. In all instances, although not clearly determined, resistance is believed to be polygenic, relatively slow to develop, and able to revert rapidly in the absence of selection pressure. Metabolic mechanisms seem to be the most important route of resistance in the arthropod systems studied to date, and in all cases, a number of biochemical and sometimes physiological mechanisms are implicated. Cross-resistance between avermectins and other insecticide classes is similarly ill-defined; however, it is apparent that there is little significant cross-resistance to avermectins except possibly in one strain of housefly (Scott, 1989). The variations in responses of field strains resistant to other insecticide classes to avermectins may in some cases be due to enhanced metabolism resulting from previous exposure to other insecticides, although there is evidence to both support and refute this supposition. A number of species have also demonstrated inherent population differences in response to avermectins without the influence of any predisposing factors. A wide variation among field-collected strains has been seen in several insects which does not constitute cross-resistance,

41

nor does it indicate that this variation predisposes insects to an increased likelihood for resistance development to avermectins. Cross-resistance between the avermectins and milbemycins (specifically abamectin, ivermectin, doramectin, and moxidectin) has been demonstrated in a number of nematode species. Because abamectin is the only avermectin compound used widely in agriculture, there have been a few studies conducted to examine cross-resistance within the class. Sensitivity to emamectin benzoate in select abamectinresistant strains has been assessed in several insects (e.g., L. trifolii, L. decemlineata, P. xylostella) and only minimal, if any, cross-resistance has been detected (Jansson and Dybas, 1998; Jansson and Rugg, unpublished data). The reasons for the lack of cross-resistance to these closely related compounds are unclear. It is possible that the changes in the derivatization of emamectin benzoate may result in the loss or protection of the chemical bonds that are subject to enzymatic cleavage in abamectin resistance. Additionally, because macrocyclic lactones have multiple sites of action and may affect a number of different receptors to activate or potentiate chloride ion influx, subtle differences in chemistry between the different avermectins and milbemycins may alter the relative activities of compounds at the different sites. This could affect the patterns and levels of cross-resistance within the class and also be a factor in the subtle differences seen in the spectra and potencies of these compounds. 5.2.6.2. Acarids

Laboratory selection of up to 15 generations of two species of tetranychid mites with abamectin resulted in no significant increase in resistance compared with a nonselected baseline colony (Hoy and Conley, 1987). Conversely, 20 rounds of selection against a predatory phytoseiid mite resulted in a gradual and modest shift in susceptibility to abamectin (Hoy and Ouyang, 1989). A recent survey of field populations of two-spotted spider mite in California (Campos et al., 1995) found that mites varied widely in their susceptibility to abamectin, with up to a 658-fold difference in susceptibility in a residual exposure assay. Levels of resistance were correlated with the amount and time of abamectin usage, and laboratory selection of a single population over 38 generations resulted in more than a 100-fold increase in resistance. Despite the higher level of resistance in a residual assay, selected mites were still considered susceptible to abamectin in a direct contact bioassay. Further, despite the wide variation in responses among field populations of mites, no field failures of the product were reported.

42 The Insecticidal Macrocyclic Lactones

In T. urticae, resistance to abamectin has been attributed to increased excretion and decreased absorption (Clark et al., 1995), monooxygenases (Campos et al., 1996), esterases (Campos et al., 1996, 1997; Jansson et al., unpublished data), glutathione-S-transferases (Clark et al., 1995), and decreased penetration (Clark et al., 1995). Levels of resistance in two spotted spider mite populations collected from the field were shown to drop markedly within 2–6 weeks after collection when maintained in the absence of selection pressure (Campos et al., 1995; Jansson et al., unpublished data). Bergh et al. (1999) reported wide variation in susceptibility to abamectin in citrus rust mite (P. oleivora) field populations with no clear relationship between susceptibility and product efficacy. While some populations from commercial groves with a history of exposure to abamectin were less susceptible to abamectin, mortality was only slightly lower than that of a pristine population. Also bioassay results indicated that the responses of these field populations did not change whether they were assessed before or after abamectin spray applications. 5.2.6.3. Helminths

Ivermectin resistance was initially detected in nematodes of sheep and goats, and was generally a result of intensive use over a number of years (Shoop, 1993). Ivermectin-resistant nematodes have been shown to have cross-resistance to other avermectins and milbemycins (Shoop et al., 1993). However, moxidectin was significantly more toxic to many parasitic nematodes (Shoop, 1993; Shoop et al., 1993; Kieran, 1994) and this difference in potency has resulted in moxidectin providing economic control of ivermectin-resistant parasites in field use situations. Conder et al. (1993) showed that the modes of action of ivermectin and moxidectin were qualitatively similar by examining changes in membrane conductance in crab muscle fibers. These workers also demonstrated cross-resistance to moxidectin in an ivermectin-resistant strain of H. contortus and confirmed that moxidectin was about fourfold more potent than ivermectin against this parasite in a jird model. Recently, Ranjan et al. (2002) reported the results of concurrent selection of H. contortus for 22 generations with ivermectin and moxidectin. These workers found that worms that had developed resistance through selection with either of these macrocyclic lactones were cross-resistant to the other compound. However, the rate of development of resistance differed between the two compounds and occurred more

slowly with moxidectin, which was also markedly more potent than ivermectin. See Pritchard (2002) for the most recent, detailed review of the status of resistance to macrocyclic lactones in nematodes. Briefly, resistance is still a major concern for parasites of sheep and has also been detected in some cattle parasites. Resistance has been found to be associated with enhanced P-glycoprotein expression resulting in increased efflux from the target site, but there are thought to be a number of possible resistance mechanisms including various amino acidgated anion subunit genes, genes associated with glutamate receptors and transporters, and genes involved in amphid structure and function. The multiplicity of possible sites for macrocyclic lactone binding and activity and their variable sensitivity to different compounds may be responsible for the differential potencies seen within and between avermectins and milbemycins and suggests that products of several genes may be involved in the mechanism of action of macrocyclic lactones. Similarly, these subtle differences in the ways in which ivermectin and moxidectin act at the molecular level could result in the differences seen between these compounds in potency, rates of development of resistance, and levels of cross-resistance.

5.2.7. Summary The macrocyclic lactones are potent insecticides, nematicides, and acaricides. The class consists of two closely related groups of compounds, the avermectins and milbemycins, which are natural fermentation products or synthetic derivatives of natural products derived from actinomycetes. The broad spectrum of activity against a variety of insects, acarids, and nematodes has led to the widespread use of these compounds in animal and human health, crop protection, and urban pest control. Ivermectin has been the most successful antiparasitic drug introduced in animal health and abamectin has been the premier agricultural miticide in recent years. Macrocyclic lactones affect invertebrates by either directly activating receptors for the neurotransmitters glutamate and GABA, or by potentiating their actions, resulting in an influx of chloride ions into nerve cells and muscle. It is currently believed that the anthelminthic properties of the avermectins are due predominantly to potentiation and/or direct opening of glutamate-gated chloride channels, whereas in insects, it is likely that avermectins bind to multiple sites, including glutamate and GABA-gated chloride channels as well as other insect chloride channels (Sattelle, 1990;

The Insecticidal Macrocyclic Lactones

Bloomquist, 1993). The chloride ion flux resulting from the opening by these compounds of chloride channels in invertebrate nerve and muscle, particularly those gated by glutamate and GABA, results in disruption of activity and loss of function in these excitable cells and accounts for the potent actions of these molecules (review: Raymond and Sattelle, 2002). Toxicity is generally greater through ingestion than by residual contact and compared with other neurotoxic agents, macrocyclic lactones are considered to be relatively slow acting. Intoxicated organisms usually die slowly, often over a period of days, and there is no quick knockdown effect. Death usually follows paralysis and immobility. The comparatively poor contact activity of these compounds plus their short environmental persistence generally results in good compatibility with beneficial and nontarget organisms. These compounds have low relative toxicities to vertebrates and are not phytotoxic. Resistance to avermectins has been demonstrated in a number of arthropods. In most instances, although not clearly determined, resistance is apparently polygenic, develops relatively slowly, and tends to revert rapidly in the absence of selection pressure. Metabolic resistance mechanisms seem to be the most important in the arthropod systems studied to date, and in all these cases a number of biochemical and physiological mechanisms are implicated. Cross-resistance between macrocyclic lactones and other insecticide classes is similarly ill-defined; however, it is apparent that there is little significant cross-resistance. The variations in responses of field strains resistant to other insecticide classes to avermectins may in some cases be due to enhanced metabolism resulting from previous exposure to other insecticides. A number of species have demonstrated wide inherent variation in response to avermectins without the influence of any predisposing factors. This natural variation in susceptibility does not constitute cross-resistance, nor is there evidence to indicate that this variation results in predisposition to develop increased resistance to avermectins. Despite the widespread, intensive use of abamectin against a number of pests with a high propensity to develop resistance, development of resistance has been relatively slow and rare. The rapid reversion of resistance in the absence of selection pressure and the lack of any confirmed cases of cross-resistance make them amenable for use in resistance management programs. However, in order to prolong the longevity of these compounds in the market place, it will be necessary to minimize selection pressure in practice, especially in high use situations, through

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proper rotation, good pest management practices, and proper resistance management strategies.

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5.3 Neonicotinoid Insecticides P Jeschke and R Nauen, Bayer CropScience AG, Monheim, Germany ß 2005, Elsevier BV. All Rights Reserved.

5.3.1. Introduction 5.3.2. Neonicotinoid History 5.3.3. Chemical Structure of Neonicotinoids 5.3.3.1. Ring Systems Containing Commercial Neonicotinoids 5.3.3.2. Neonicotinoids Having Noncyclic Structures 5.3.3.3. Bioisosteric Segments of Neonicotinoids 5.3.3.4. Physicochemistry of Neonicotinoids 5.3.3.5. Proneonicotinoids 5.3.4. Biological Activity and Agricultural Uses 5.3.4.1. Efficacy on Target Pests 5.3.4.2. Agricultural Uses 5.3.4.3. Foliar Application 5.3.4.4. Soil Application and Seed Treatment 5.3.5. Mode of Action 5.3.6. Interactions of Neonicotinoids with the Nicotinic Acetylcholine Receptor 5.3.6.1. Selectivity for Insect over Vertebrate nAChRs 5.3.6.2. Whole Cell Voltage Clamp of Native Neuron Preparations 5.3.6.3. Correlation between Electrophysiology and Radioligand Binding Studies 5.3.7. Pharmacokinetics and Metabolism 5.3.7.1. Metabolic Pathways of Commercial Neonicotinoids 5.3.8. Pharmacology and Toxicology 5.3.8.1. Safety Profile 5.3.9. Resistance 5.3.9.1. Activity on Resistant Insect Species 5.3.9.2. Mechanisms of Resistance 5.3.9.3. Resistance Management 5.3.10. Applications in Nonagricultural Fields 5.3.10.1. Imidacloprid as a Veterinary Medicinal Product 5.3.11. Concluding Remarks and Prospects

53 54 56 57 62 64 67 70 72 72 73 75 75 75 76 76 76 78 79 79 82 83 87 87 89 91 91 92 95

5.3.1. Introduction p0005

The discovery of neonicotinoids as important novel pesticides can be considered a milestone in insecticide research of the past three decades. Neonicotinoids represent the fastest-growing class of insecticides introduced to the market since the commercialization of pyrethroids (Nauen and Bretschneider, 2002). Like the naturally occurring nicotine, all neonicotinoids act on the insect central nervous system (CNS) as agonists of the postsynaptic nicotinic acetylcholine receptors (nAChRs) (Bai et al., 1991; Liu and Casida, 1993a; Yamamoto, 1996; Chao et al., 1997; Zhang et al., 2000; Nauen et al., 2001), but, with remarkable selectivity and efficacy against pest insects while being safe for mammals. As a result of this mode of action there is no cross-resistance to conventional insecticide classes, and therefore the neonicotinoids have begun replacing pyrethroids, chlorinated hydrocarbons,

organophosphates, carbamates, and several other classes of compounds as insecticides to control insect pests on major crops (Denholm et al., 2002). Today the class of neonicotinoids are part of a single mode of action group as defined by the Insecticide Resistance Action Committee (IRAC; an Expert Committee of Crop Life) for pest management purposes (Nauen et al., 2001). Neonicotinoids are potent broad-spectrum insecticides possessing contact, stomach, and systemic activity. They are especially active on hemipteran pest species, such as aphids, whiteflies, and planthoppers, but they are also commercialized to control coleopteran and lepidopteran pest species (Elbert et al., 1991, 1998). Because of their physicochemical properties they are useful for a wide range of different application techniques, including foliar, seed treatment, soil drench, and stem application in

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several crops. Due to the favorable mammalian safety characteristics (Matsuda et al., 1998; Yamamoto et al., 1998; Tomizawa et al., 2000) neonicotinoids like imidacloprid are also important for the control of subterranean pests, and for veterinary use (Mencke and Jeschke, 2002).

5.3.2. Neonicotinoid History

p0020

In the early 1970s, the former Shell Development Company’s Biological Research Center in Modesto, California, invented a new class of nitromethylene heterocyclic compounds capable of acting on the nAChR. Starting with a random screening to discover lead structures from university sources, Shell detected the 2-(dibromo-nitromethyl)-3-methyl pyridine (SD-031588) (Figure 1) from a pool of chemicals from Prof. Henry Feuer of Purdue University (Feuer and Lawrence, 1969), which revealed an unexpected low-level insecticidal activity against housefly and pea aphid. Further structural optimization of this insecticide lead structure led to the active six-membered tetrahydro-2-(nitromethylene)-2H-1,3-thiazine, nithiazine (SD-03565, SKI-71) (Soloway et al., 1978, 1979; Schroeder and Flattum, 1984; Kollmeyer et al., 1999). The molecular design by Shell appears rational and straightforward. The chemistry had been largely concentrated on the nitromethylene enamine skeleton (Kagabu, 2003a). Today, this early prototype can be considered as the first generation of the socalled neonicotinoid insecticides (Figure 2). Nithiazine showed higher activity than parathion against housefly adults (Musca domestica), and 1662 times higher activity against the target insect, the lepidopteran corn earworm larvae (Helicoverpa zea), combined with good systemic behavior in plants and low mammalian toxicity (Soloway et al., 1978, 1979; Kollmeyer et al., 1999; Tomizawa and Casida,

2003). However, due to the photochemically unstable 2-nitromethylene chromophore (Figure 3) in the field tests, nithiazine was never commercialized for broad agricultural use (Soloway et al., 1978, 1979; Kagabu and Medej, 1995; Kagabu, 1997a; Kollmeyer et al., 1999). Alternatively, photostabilization using the formyl moiety was not adequate for practical application (Kollmeyer et al., 1999). Nevertheless, a knock-down fly product against M. domestica containing nithiazine as the active ingredient of a housefly trap device for poultry and animal husbandry has recently been commercialized (Kollmeyer et al., 1999). In the early 1980s synthesis work was initiated at Nihon Tokushu Noyaku Seizo K. K. (presently Bayer CropScience K. K.) on the basis of this first remarkable neonicotinoid lead structure and the unique insecticidal spectrum of activity (Kagabu et al., 1992; Moriya et al., 1992). Instead of the lepidopteran larva H. zea, the target insect for studying structure–activity relationships (SARs) and optimizing biological activity was the green rice leafhopper (Nephotettix cincticeps), because it is a major hemipteran pest of rice in Japan (Kagabu, 1997a). At the beginning of the project, a new pesticide screening method using rice seedlings was developed for continuous monitoring of the combined systemic and contact activities of compounds over 2 weeks against leafhoppers and planthoppers (Sone et al., 1995). The six-membered 2-(nitromethylene)-tetrahydro1,3-thiazine ring was replaced with different N1substituted N-heterocyclic ring systems. Starting with 1-methyl-2-(nitromethylene)-imidazolidine, it was found that the activity depends on ring size (5 > 6 > 7-ring system). The N1-benzylated 2(nitromethylene)-imidazolidine 5-ring was the most active one. Introduction of substituted benzyl residues in the 1-position, like the para-chlorobenzyl moiety, and of nitrogen-containing N-hetarylmethyl

Figure 1 Development of nithiazine (SKI-71) by Shell. (Data from Kollmeyer, W.D., Flattum, R.F., Foster, J.P., Powel, J.E., Schroeder, M.E., et al., 1999. Discovery of the nitromethylene heterocycle insecticides. In: Yamamoto, I., Casida, J.E. (Eds.), Neonicotinoid Insecticides and the Nicotinic Acetylcholine Receptor. Springer, New York, pp. 71–89 and Kagabu, S., 2003a. Molecular design of neonicotinoids: past, present and future. In: Voss, G., Ramos, G. (Eds.), Chemistry of Crop Protection: Progress and Prospects in Science and Regulation. Wiley–VCH, New York, pp. 193–212.)

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f0010

55

Figure 2 Chemical structures of the first generation neonicotinoid nithiazine and commercialized neonicotinoids displaying the different types of pharmacophors. CPM, 6-chloro-pyrid-3-ylmethyl; CTM, 2-chloro-1,3-thiazol-5-ylmethyl; TFM, (
)-6-tetrahydro-fur3-ylmethyl. (Reproduced with permission from Jeschke, P., Schindler, M., Beck, M., 2002. Neonicotinoid insecticides: retrospective consideration and prospects. Proc. BCPC: Pests and Diseases 1, 137–144.)

Figure 3 Electronic absorption (lmax in nm), water solubility (g l1 at 20  C), and insecticidal efficacy (ppm) of nithiazine (SKI-71), nitromethylene (NTN32692), and imidacloprid (NTN33893) against green rice leafhopper (Nephotettix cincticeps). LC90, concentration at which >90% of insects are killed (Mencke and Jeschke, 2002).

residues, like the pyrid-3-ylmethyl moieties, enhanced the insecticidal activity by a factor of 5 and 25 when compared with the N1-benzylated 2-(nitromethylene)-imidazolidine (Kagabu, 1997a). Yamamoto had previously recognized that the pyrid3-ylmethyl residue was an essential moiety for insecticidal activity, by referring to the nAChR agonist model of Beers and Reich (1970), and thus synthesized a number of pyrid-3-yl-amines (Yamamoto and Tomizawa, 1993).

This chemical optimization procedure led to discovery of 1-(6-chloro-pyrid-3-ylmethyl)-2-nitromethylene-imidazolidine (NTN32692), which had over 100-fold higher activity than nithiazine against N. cincticeps (using a strain resistant to organophosphorus compounds, methylcarbamates, and pyrethroides) (Kagabu et al., 1992; Moriya et al., 1992). However, the 2-nitromethylene chromophore of NTN32692 absorbs strongly (Figure 3) in sunlight (wavelength 290–400 nm), and rapidly decomposes

p0035

56 Neonicotinoid Insecticides

Figure 4 Structural segments for neonicotinoids.

in the field to noninsecticidal compounds. In order to avoid absorption of sunlight, a more stable functional group was necessary. After preparation of about 2000 compounds, imidacloprid (NTN33893) containing a 2-(N-nitroimino) group (see Section 5.3.3.1.1), the first member of the secondgeneration neonicotinoids, emerged from this project (Elbert et al., 1990, 1991; Bai et al., 1991; Nauen et al., 2001). The presence of an N-nitroimino group at the 2-position of the imidazolidine ring makes little difference to insecticidal activity compared with compounds that contain a nitomethylene group at this position, but the presence of ›N NO2 w significantly reduces affinity for the receptor (Liu et al., 1993b; Tomizawa and Yamamoto, 1993; Yamamoto et al., 1998). This suggests that the reduction in binding, resulting from the presence of the nitrogen atom at the 2-position, is compensated by the increase in hydrophobicity, which enhances transport to the target sites (Yamamoto et al., 1998; Matsuda et al., 2001). Compared with nithiazine, the biological efficacy of imidacloprid against the green rice leafhopper could be enhanced 125-fold. Furthermore, imidacloprid is about 10 000-fold more insecticidal than (S)-nicotine, a natural insecticide (Tomizawa and Yamamoto, 1993; Yamamoto et al., 1995). This breakthrough to the novel systemic insecticide imidacloprid was achieved by coupling a special heterocyclic group, the 6-chloro-pyrid-3ylmethyl (CPM) residue (Figure 4), to the 2-(Nnitroimino)-imidazolidine building-block, new in this class of chemistry. With the introduction of the insecticide imidacloprid to the market in 1991 Bayer AG began a successful era of the so-called CNITMs (chloronicotinyl

insecticides syn. neonicotinoids), a milestone in insecticide research. Imidacloprid has, in the last decade, become the most successful, highly effective, and best selling insecticide worldwide used for crop protection and veterinary pest control. In connection with these excellent results, a parallel change to other electron-withdrawing substituents like 2-N-cyanoimino, having essentially shorter maximum electronic absorptions than 290 nm, led to the discovery of thiacloprid (see Section 5.3.3.1.2), a second member of the CNI group. Attracted by Bayer’s success with imidacloprid, several different companies such as Takeda (now Sumitomo Chemical Takeda Agro), Agro Kanesho, Nippon Soda, Mitsui Toatsu (now Mitsui Chemicals, Inc.), Ciba Geigy (now Syngenta), and others initiated intensive research and developed their own neonicotinoid insecticides. Research in these companies was facilitated because neonicotinoid chemistry showed a relatively broad spectrum of activity (Wollweber and Tietjen, 1999; Roslavtseva, 2000). Since the market introduction of imidacloprid, neonicotinoids have become the fastest-growing class of chemical insecticides. This tremendous success can be explained by their unique chemical and biological properties, such as broad-spectrum insecticidal activity, low application rates, excellent systemic characteristics such as uptake and translocation in plants, new mode of action, and favorable safety profile.

5.3.3. Chemical Structure of Neonicotinoids In general, all these commercialized or developed compounds can be divided into ring systems

p0045

Neonicotinoid Insecticides

containing neonicotinoids, and neonicotinoids having noncyclic structures which differ in their molecular characteristics. Structural requirements for both ring systems containing neonicotinoids and neonicotinoids having noncyclic structures consist of different segments (Nauen et al., 2001): the bridging fragment [R1 R2] (i) and for the noncyclic type w compounds, the separate substituents [R1, R2] (i), the heterocyclic group [Het] (ii), the bridging chain [ CHR ] and the functional group [›X Y] w w w as part of the pharmacophore [ N C(E)›X Y] w w w (iii) (Figures 2 and 4, Table 1). The methylene group is normally used as the bridging chain ( CHR with R ¼ H). Other groups w w such as an ethylene or substituted methylene group decrease the biological activity. The pharmacophore (iii) can be represented by the group [ N C(E) w w ›X Y], where ›X Y is an electron-withdrawing w w 0 group and E is a NH, NR , CH2, O, or S moiety. It is well known that the pharmacophore type influences the insecticidal activity of the neonicotinoids. In general, the greatest insecticidal activity is observed for the compounds containing the nitroenamine (nitromethylene) [ N C(E)›CH NO2; E ¼ S, N], w w w nitroguanidine [ N C(E)›N NO2; E ¼ N], or w w w cyanoamidine [ N C(E)›N CN; E ¼ S, CH3] w w w pharmacophore (Shiokawa et al., 1995). Besides its influence on biological activity, the pharmacophore is also responsible for some specific properties such as photolytic stability, degradation in soil, metabolism in plants, and toxicity to different animals (Kagabu et al., 1992; Moriya et al., 1992; Minamida et al., 1993; Tomizawa and Yamamoto, 1993; Shiokawa et al., 1994; Tabuchi et al., 1994). The term ‘‘neonicotinoid’’ (Yamamoto, 1998, 1999) was originally proposed for imidacloprid and related insecticidal compounds with structural similarity to the insecticidal alkaloid (S)-nicotine, which has a similar mode of action (Tomizawa and Yamamoto, 1993; Tomizawa and Casida, 2003). In addition, a variety of terms have been used to subdivide this chemical class based on structural fragments (Figure 4): (a) chloronicotinyls (CNIs, heterocyclic group CPM) (Leicht, 1993), (b) thianicotinyls (heterocyclic group CTM) (Maienfisch et al., 1999a), (c) furanicotinyls (heteroalicyclic group, TFM) (Wakita et al., 2003), (d) nitroimines or nitroguanidines, (e) nitromethylenes, (f) cyanoimines (functional group as part of the pharmacophore), (g) first generation with moiety CPM, (h) second generation with moiety CTM or TFM (Maienfisch et al., 1999a, 2001b), and third generation with moiety TFM (regarding the type of the heterocyclic group) (Wakita et al., 2003).

57

5.3.3.1. Ring Systems Containing Commercial Neonicotinoids

Ring systems containing neonicotinoids can have as bridging fragment [R1 R2] (i) an alkylene group in w the building-block, e.g., ethylene for imidacloprid and thiacloprid (five-membered ring systems) which can also be interrupted by heteroatoms like oxygen (six-membered ring systems) as shown in the case of thiamethoxam. 5.3.3.1.1. 1-[(6-Chloro-3-pyridinyl)-methyl]-Nnitro-2-imidazolidinimine (imidacloprid, NTN 33893) Imidacloprid was discovered in 1985 as a novel insecticide with a unique structure and with hitherto unrecognized insecticidal performance (Moriya et al., 1992, 1993; Kagabu, 1997a, 2003a). The trade names for soil application/seed treatment are AdmireTM and GauchoÕ and the trade names for foliar application are ConfidorÕ and ProvadoÕ, while for termite control it is PremiseÕ (see Section 5.3.10). The first laboratory synthesis was carried out by reduction of 6-chloronicotinoyl chloride to 2-chloro-5-hydroxymethyl-pyridine using an excess NaBH4 in water, and conversion to the chloride by SOCl2. Imidacloprid was obtained by the coupling reaction of 2-chloro-5-chloromethyl-pyridine (CCMP) (Diehr et al., 1991; Wollweber and Tietjen, 1999) with the 5-ring building-block 2-nitro-imino-imidazolidine, in acetonitrile with potassium carbonate as base. This method was also successfully applied to the synthesis of [3H]imidacloprid using NaB[3H]4 (Latli et al., 1996). Differences in photostability between the 2-nitromethylene group in compound NTN32692 (lmax ¼ 323 nm) and the 2-(N-nitroimino) group in imidacloprid (lmax ¼ 269 nm) (Figure 3) were examined by quantum chemical methods like AM1 and ab initio calculations (Kagabu, 1997a; Kagabu and Akagi, 1997). Crystallographic analysis of imidacloprid revealed a coplanar relationship of the imidazolidine 5-ring to the nitroimino group in position 2 (Born, 1991; Kagabu and Matsuno, 1997). Shorter ring C N bond lengths and longer exocyclic C›N w bond lengths were observed, suggesting the formation of a planar electron delocalized diene system, a so-called push–pull olefin (Kagabu and Matsuno, 1997). An intramolecular hydrogen bond between 1 NH  O2N N›C2 was confirmed by characterisw tic resonances in the nuclear magnetic resonance (NMR) spectra (correlation spectroscopy (COSY), nuclear overhauser enhancement spectroscopy (NOESY)). In addition, the infrared (IR) spectrum showed a highly chelated 1NH absorption by intense bands at 3356 and 3310 cm1 (Kagabu et al., 1998).

p0075

p0080

Table 1 Chemical classification of nithiazine and commercialized neonicotinoids Scientific name Development codes

Empirical formula

Molecular weight (g mol 1)

C5H8N2O2S

160.2

C9H10ClN5O2

255.7

Common name

IUPAC name

Chemical Abstracts name

CAS registry no.

Nithiazine

2-Nitromethylene-1,3-thiazinane

Tetrahydro-2-(nitromethylen)-2H-1,3-thiazine

[58842-20-9]

Imidacloprid

1-(6-Chloro-3-pyridylmethyl)-Nnitroimidazolidin-2-ylidenamine N-[3-(6-Chloro-pyridin-3-ylmethyl)-thiazolidin-2ylidene]-cyanamide 3-(2-Chloro-thiazol-5-ylmethyl)-5-methyl-1,3,5oxadiazinan-4-ylidene (nitro)amine (E )-N-(6-Chloro-3-pyridylmethyl)-N-ethyl-N 0 methyl-2-nitrovinylidenediamine

1-[(6-Chloro-3-pyridinyl)-methyl]-N-nitro-2imidazolidinimine Cyanamidine, [3-[(6-chloro-3-pyridinyl)methyl-2thiazolidinylidene] 3-[(2-Chloro-5-thiazolyl)methyl]tetrahydro-5methyl-N-nitro-4H-1,3,5-oxadiazin-4-imine N-[(6-Chloro-3-pyridinyl)methyl]-N-ethyl-N 0 -methyl2-nitro-1,1-ethenediamine

[138261-41-3]

SD-035651a IN-A0159b NTN 33893

[111988-49-9]

YRC 2894

C10H9ClN4S

252.8

[153719-23-4]

CGA2930 343

C8H10ClN5O3S

291.7

TI-304

C11H15ClN4O2

270.7

(E )-N 1-[(6-Chloro-3-pyridyl)methyl]-N 2-cyanoN 1-methyl-acetamidine (E )-1-(2-Chloro-1,3-thiazol-5-ylmethyl)-3methyl-2-nitroguanidine (RS )-1-methyl-2-nitro-3-(tetrahydro-3furylmethyl)guanidine

(E )-N-[(6-chloro-3-pyridinyl)methyl]-N 0 -cyano-Nmethyl-ethanimidamide [C(E )]-N-[(2-Chloro-5-thiazolyl)methyl]-N 0 -methylN 00 -nitroguanidine N-Methyl-N 0 -nitro-N 00 -[(tetrahydro-3furanyl)methyl]guanidine

[120738-89-8] [150824-47-8] (E-isomer) [135410-20-7]

NI-25

C10H11ClN4

222.7

[210880-92-5]

TI-435

C6H8ClN5O2S

249.7

[165252-70-0]

MTI 446

C7H14N4O3

202.2

Thiacloprid Thiamethoxam Nitenpyram

Acetamiprid Clothianidin (
)-Dinotefuran

a

Shell/DuPont. DuPont.

b

Neonicotinoid Insecticides

Studies were also done using comparative molecular field analysis (CoMFA), a technique for analysis of three-dimensional quantitative structure–activity relationships (3D QSAR) (Cramer et al., 1988; Okazawa et al., 1998). The model showed that the nitrogen atom of the imidacloprid CPM residue interacts with a hydrogen-donating site of nAChR, and the nitrogen atom at the 1-position of the imidazolidine 5-ring interacts with a negatively charged domain (Okazawa et al., 2000). In comparison with (S)nicotine the selective insecticidal mode of action of imidacloprid was well examined in detail by different researchers (Yamamoto et al., 1995; Kagabu, 1997b; Matsuda et al., 1999; Tomizawa, 2000). There studies focused on the chemical structure, state of ionization, and hydrophobicity of both molecules (Figure 5). The principal target insect pests of imidacloprid in agricultural and veterinary medicinal use are sucking insects. Since imidacloprid provides a partial positively charged dþ nitrogen atom (not ionized, full or partial) instead of an ammonium head as does (S)-nicotine, imidacloprid has little restriction for translocation into the target area compared

59

to (S)-nicotine (Graton et al., 2003). The interatomic distances between the 2-(N-nitroimino)-imidazolidine nitrogen and the pyridyl nitrogen are 5.9 A˚, which is a suitable value to bind to the electron-rich and hydrogen-donating recognition sites on nAChR (Yamamoto et al., 1995; Matsuda et al., 1999). The electron-donating atom for hydrogen bonding at 5.9 A˚ apart from the fully or partial positive charged dþ nitrogen is required for potent interaction with the anionic subsite of the insect nAChR (Yamamoto et al., 1995; Tomizawa, 2000). In mammals, imidacloprid has weak effects on any nAChR, while nicotine affects mostly peripheral nACRs resulting in higher toxicity. The deduced electron deficiency of the nitrogen atom of imidacloprid was proved explicitly by 15 N NMR spectroscopic measurements (Yamamoto et al., 1995). Tomizawa et al. (2000) recently claimed that the N-nitro group of imidacloprid is much more important than the bridgehead nitrogen in electrostatic interaction with nAChRs’ because a Mulliken charge of the bridgehead nitrogen, calculated by the MNDO method combined with the PM3 method (semi-empirical molecular orbital technique

Figure 5 Insecticidal mode of action of (S)-nicotine and imidacloprid. (Adapted from Tomizawa, M., 2000. Insect nicotinic acetylcholine receptors: mode of action of insecticide and functional architecture of the receptor. Jap. J. Appl. Entomol. Zool. 44, 1–15.)

60 Neonicotinoid Insecticides

for calculating electronic structure), was only marginally positive. However, Matsuda et al. (2001) suggested, that the electron-withdrawing power of the N-nitro group might be increased by a hydrogen bond involving the N-nitro group oxygen and hydrogen-bondable and positively charged amino acid residue, namely lysine or arginine, on the insect receptor, thereby strengthening its interaction with imidacloprid. Kagabu (1996) predicted the important contribution of this N-nitro group and its hydrogen-bondable property for insecticidal activity. 5.3.3.1.2. [3-(6-Chloro-3-pyridinyl)methyl-2-thiazolidinylidene]-cyanamidine (thiacloprid, YRC 2894) The second member of the CNITM family from Bayer AG launched in 2000 was thiacloprid (Elbert et al., 2000; Yaguchi and Sato, 2001). Similar to imidacloprid, this neonicotinoid thiacloprid (YRC 2894, registered worldwide under the trade name CalypsoTM, and in Japan and Switzerland under the trade names BariardTM and AlantoTM), also contains the unique CPM moiety attached to the cyclic 2-(cyanoimino)-thiazolidine (CIT) buildingblock. Thiacloprid can be synthesized by a convergent one-step technical process starting from two key intermediates, CIT and CCMP (Diehr et al., 1991; Wollweber and Tietjen, 1999; Jeschke et al., 2001). Thiacloprid crystallizes in two different modifications depending on the solvent. The compound crystallizes from dichloromethane as form I (melting point 136  C) and from isopropanol as form II (melting point 128.3  C). From its physicochemical data, the technical ingredient is form I.

The results of X-ray crystallographic analysis and an X-ray powder diffraction profile indicate that each form, I (0.6  0.3  0.2 mm3) and II (0.5  0.2  0.1 mm3), crystallizes with different cell constants in a monoclinic crystal system in the space group P21/c (Jeschke et al., 2001). Figure 6 shows the Ortep plot of the crystal structure of the form I. Interestingly, in the crystal lattice of form II, two symmetric independent molecules in the asymmetric unit cell are fixed (Figure 6). With regard to the configuration of the pharmacophore [ N C(S)›N CN] moiety, thiacloprid w w w exists only in the Z-configuration in both forms, I and II. In conjunction with the X-ray results, thiacloprid exists in solution exclusively in the stable Z-configuration. With respect to the C›N double bond, only one configuration is seen in all NMR spectra (1H, 13C, HMQC, HMBC, NMR spectroscopy), deduced from very sharp proton and carbon signals. In conjunction with X-ray results, there is no evidence for isomerization of the Z-isomer in DMSO-d6 or CDCl3 solution. Thiacloprid shows a single peak maximum at 242 nm in its ultraviolet (UV) spectrum, and it does not absorb natural sunlight. This explains thiacloprid’s better photostability in comparison to other neonicotinoids (Jeschke et al., 2001). Starting from forcefield methods (MMFF94s), the final energies, geometries, and properties were obtained at DFT (Becke and Perdew (BP) functional, triple-zeta valence plus (TZVP) polarization basis, conductor-like screening model – realistic solvent (COSMO-RS) for solvent effects; Perdew, 1986;

Figure 6 Ortep plot of thiacloprid form I and form II (Jeschke et al., 2001).

p0105

Neonicotinoid Insecticides

61

Figure 7 DFT (BR/TZVP) optimized geometry of thiacloprid (Jeschke et al., 2001). (a) Atomic charges projected onto a Conolly surface; (b) some interatomic distances in A˚.

Becke, 1988; Klamt, 1995) level of theory (Jeschke et al., 2001) (Figure 7). It was found that, in the gas phase, Z-configured thiacloprid is about 4 kcal mol1 lower in energy than the E-isomer. In water, this difference is increased by approximately 1 kcal mol1. The preference for the Z-configuration stems mainly from steric reasons. The nitrile moiety of the Z-isomer has virtually no close contacts to any other atom (sulfur to nitrile-carbon ˚ ). In the E-isomer, however, the largest distance 3.1 A possible distance, between the nitrile-carbon and the hydrogen atoms attached to the bridging carbon, is around 2.3 A˚ . Quantum chemical calculations show that the strong delocalization of the C›N double bond does not reduce the ‘‘double bond character’’ significantly (i.e., the torsional barrier for rotation around the C›N double bond is not decreased). The molecular dipole moment (derived from the electric field dependence of the DFT wave function) is calculated to be 10.2 D in water. This, together with the spatial relation of the pharmacophoric feature ( N C(S)›N CN), is very much in line with the w w w pharmacophore model of imidacloprid. 5.3.3.1.3. 3-[(2-Chloro-5-thiazolyl)methyl]tetrahydro-5-methyl-N-nitro-4H-1,3,5-oxadiazin-4-imine (thiamethoxam, CGA 2930 343) Ciba started a research program on neonicotinoids in 1985 (Maienfisch et al., 2001a), and investigated variations of the imidacloprid nitroimino-heterocycle (Maienfisch et al., 2001b). Thiamethoxam (Senn et al., 1998; Maienfisch et al., 1999a, 1999b) was discovered, and developed by Ciba Crop Protection (from 1996 Novartis Crop Protection, now Syngenta Crop Protection) in 1991. Thiamethoxam has been marketed since 1998 under the trademarks ActaraÕ

for foliar and soil treatment, and CruiserÕ for seed treatment. Thiamethoxam can be synthesized starting from N-methyl-nitroguanidine by treatment with formaldehyde in the presence of formic acid (Go¨ bel et al., 1999). Finally alkylation with 2-chlorothiazol-5-ylmethylchloride in N,N-dimethyl-formamide and potassium carbonate as a base afforded the active ingredient in good yields (Maienfisch et al., 1999a). The SAR profile of this compound demonstrates the rather limited variability of the pharmacophore ( N C(N)›N NO2). Insecticidal activity was only w w w observed if the functional group is a strongly electronwithdrawing group and has a hydrogen accepting head, like N-nitroimino and N-cyanoimino. As pharmacophore N-substituent, a methyl group is clearly superior to hydrogen, an acyl substituent or C2 C4 alkyl group. The biological results have led w to a general SAR profile for these novel compounds, which can be summarized as follows (Maienfisch et al., 1999b, 2002): 1. All variations of the pharmacophore (N C(N)›N NO2) resulted in a loss of activity. w w 2. Substitution of the CTM moiety by another Ncontaining heterocycle reduced the overall insecticidal activity, whereas CPM gave the best results. 3. A perhydro-1,3,5-oxadiazine ring system is clearly better than all other heterocyclic systems like perhydro-1,3,5-thiadiazines, hexahydro-1,3,5triazines, or hexahydro-1,3-diazines. 4. Introduction of N-methyl at the 5-position led to strong increase of insecticidal activity (in contrast to the SAR of imidacloprid), whereas other substituents resulted in clear loss of activity.

62 Neonicotinoid Insecticides

5.3.3.2. Neonicotinoids Having Noncyclic Structures

From the new chemical class of ring system containing neonicotinoids noncyclic structures having the same mode of action can also be deduced. These noncyclic type neonicotinoids can have as separate substituents [R1, R2] (i), e.g., for R1 hydrogen or alkyl like methyl (acetamiprid) and ethyl (nitenpyram) and in the case of E ¼ NH for the substituent R2 alkyl like methyl (clothianidin and dinotefuran), respectively. 5.3.3.2.1. N-[(6-chloro-3-pyridinyl)methyl]-N-ethylN0 -methyl-2-nitro-1,1,-ethenediamine (nitenpyram, TI-304) Starting from the cyclic nithiazine (Soloway et al., 1978), the acyclic nitenpyram (BestguardTM) was discovered during optimization of substituents of an acyclic nitroethene (Minamida et al., 1993). The insecticidal activity of 1-(3-pyridylamino)-2-nitroethene compounds against the brown planthopper and the green leafhopper were studied. It was found that the 1-methylamino-1(pyrid-3-ylmethylamino)-2-nitroethene was more active than the corresponding 1-ethyl, 1-methoxy, or 1-thiomethoxy derivatives. Introduction of sterically large amino groups like ethylamino, isopropylamino, hydrazino, N-pyrrolidino, or Nmorpholino decreased the activity against the green rice leafhopper. Furthermore, the methylene bridge between the pyrid-3-yl and nitrogen was replaced with other linkages. However, all attempts at shortening and lengthening the linkage failed to increase the insecticidal activities against the brown planthopper and the green rice leafhopper (Akayama and Minamida, 1999). Heterocyclic aromatic substituents, different from pyrid-3-yl, were incorporated into the nitroethen structure. The replacement of pyrid-3-yl with 6-fluoro-pyrid3-yl, 6-chloro-pyrid-3-yl, 6-bromo-pyrid-3-yl, and 2-chloro-1,3-thiazol-5-yl enhanced the activity against the brown planthopper. 5.3.3.2.2. (E)-N-[(6-chloro-3-pyridinyl)methyl]N0 -cyano-N-methyl-ethanimidamide (acetamiprid, NI-25) Acetamiprid (Takahashi et al., 1992; Matsuda and Takahashi, 1996) has an N-cyanoamidine structure, which contains in analogy to imidacloprid and thiacloprid the CPM moiety. This neonicotinoid was invented during a search for nitromethylene derivatives by Nippon Soda Co. Ltd. in 1989 and was registered in 1995 in Japan. The insecticide is marketed under the trade name MospilanÕ for crop protection. The noncyclic acetamiprid was discovered by optimization studies of special 2-N-cyanoimino compounds with an imidazolidine 5-ring

obtained from Nihon Bayer (Yamada et al., 1999). The noncyclic 2-nitromethylene and 2-N-nitroimine derivatives are less active than the corresponding ring structures, but numerous noncyclic derivatives show excellent activities against the armyworm and aphid as well as the cockroach. Regarding the substituent on the amino group, the N-methyl group exhibited the highest activity against the diamondback moth, while derivatives with hydrogen, Nmethyl, and N-ethyl showed potent activity against the cotton aphid (Yamada et al., 1999). 5.3.3.2.3. [C(E)]-N-[(2-chloro-5-thiazolyl)methyl]N0 -methyl-N00 -nitroguanidine (clothianidin, TI-435) As a result of continuous investigations of noncyclic neonicotinoids, researchers from Nihon Bayer Agrochem and Takeda in Japan found that these noncyclic neonicotinoids showed high activities against sucking insects. In the optimization process, Takeda researchers were able to demonstrate that compounds containing the nitroguanidine moiety, coupled with the thiazol-5-ylmethyl residue, have increased activity against some lepidopteran pests (Uneme et al., 1999). This is in accordance with the general pharmacophore [ N C(E)›X Y], where › w w w X Y is an electron-withdrawing group such as › w N NO2, and E represents the NH Me unit. After w w further optimization in this subclass, clothianidin (TI-435) emerged as the most promising derivative from this program (Ohkawara et al., 2002; Jeschke et al., 2003). Clothianidin has already been registered in Japan for foliar and soil applications under the trade names DantotsuTM and FullswingTM (Sumitomo Chemical Takeda Agro), and also in Korea, Taiwan, and other countries. Registrations have also been granted in North America and Europe for seed treatment under the brand name PonchoÕ (Bayer CropScience). In the novel noncyclic structure of clothianidin (TI-435) (i.e., R ¼ H; E ¼ NHMe), the N-nitroguanidine pharmacophore [ N C(N)›N NO2] is w w w similar to that of imidacloprid, but the CPM group has been replaced by the CTM moiety (Ohkawara et al., 2002) (Figure 2). Clothianidin crystallizes under normal laboratory conditions using common solvents as needlelike crystals containing no additional solvent or water molecules. The best-quality crystals for X-ray structure analysis were obtained by slow evaporation of a methanol/H2O (1 : 1) solution at room temperature. Figure 8 shows a photograph using polarized light of the crystal needle with the dimensions 0.90  0.30  0.07 mm3 used for Xray structure analysis (Jeschke et al., 2003). Clothianidin (TI-435) was subjected to conformational sampling using forcefield methods (MMFF94s).

s0055 p0135

Neonicotinoid Insecticides

The calculated free energies indicate that at room temperature (RT), the preferred orientation of the N-nitroimino group is in the trans position; the Z-isomer with lowest energy is more than 2.6 kcal mol1 above the optimal E-isomer. NMR experiments are in agreement that the N-nitroimino group strongly prefers one orientation only. From calculations as well as the X-ray structure, one can see that the three C N bonds involving atom C5 w have some double bond character. It is worth mentioning that the C5 N3 bond, which is the only w formal C›N double bond within the N-nitroimino moiety, is slightly longer than the formal single bonds C5 N5 and C5 N2. This is reflected by the w w torsional angles found around the C N bonds durw ing conformational analysis; the respective values   are all 180 or 180 . The N-methyl group can flip easily from the anti position into a syn position. The energies of the respective clothianidin (TI-435) conformers, relative to the optimal structure, are below 1.5 kcal mol1. All these findings are in line with the experimental 13C NMR spectrum, which shows a relatively broad singlet at 138.8 ppm for the atom C3. This can already be understood quali-

Figure 8 Digital stereo photomicrograph of a single crystal of clothianidin (TI-435) grown from methanol/water at room temperature (Jeschke et al., 2003).

63

tatively from the conformational arguments, like rotations of the C N single bonds and modificaw tions of the positioning of the CTM moiety (Jeschke et al., 2003). 5.3.3.2.4. (
)-N-methyl-N0 -nitro-N00 -[(tetrahydro3-furanyl)methyl]guanidine (dinotefuran, MTI446) (
)-Dinotefuran (MTI-446) was discovered by Mitsui Chemicals Inc., and is highly effective as an agonist of nAChRs (Zhang et al., 2000). It has a broad spectrum of activity against insect pests and has low mammalian toxicity (Kodaka et al., 1998, 1999a, 1999b; Hirase, 2003). The trade names for the commercialized product are StarkleÕ and AlbarinÕ. Similar to imidacloprid and clothianidin, it has a N-nitroguanidine pharmacophore [ N C(N)›N NO2], but an alicyclic (
)-tetrahyw w w dro-3-furylmethyl (TFM) moiety instead of the halogenated heteroaromatic CPM or CTM moiety in the other neonicotinoids. The discovery of (
)-dinotefuran resulted from the idea of incorporating an N-nitroimino fragment into the structure of acetylcholine (Kodaka et al., 1998; Wakita et al., 2003). (
)-Dinotefuran bears a nonaromatic oxygen atom in the position corresponding to that of the aromatic nitrogen atom of the other neonicotinoids. The potencies of (
)-dinotefuran and a competitive nAChR antagonist (a-bungarotoxin, BGT) in inhibiting [3H]epibatidine binding to Periplaneta americana nerve cord membranes were examined (Mori et al., 2001). (
) and (þ) Dinotefuran inhibited [3H]epibatidine binding with IC50 (inhibitory concentration, where 50% of the tritiated ligand is displaced in membrane preparations from cockroach nerve) values of 890 and 856 nM, respectively. The ()-enantiomer was about twofold less effective. In contrast the (þ)-enantiomer was approximately 50-fold more insecticidal than the ()-enantiomer of dinotefuran (Table 2). The SAR between the tetrahydrofuran ring moiety and the insecticidal activity was investigated (Kiriyama and Nishimura, 2002), and it was found that:

Table 2 Potency of dinotefuran and its isomers in inhibiting [3H]EPI and [3H]a-BGT binding, and their in vivo activities

Compound

[ 3H]EPI binding, IC50 (nM)a

[ 3H]a-BGT binding, IC50 (nM)a

Knockdown, KD50 (nmol g1)

Insecticidal activity, LD50 (nmol g1)a

(
)-Dinotefuran (þ)-Dinotefuran ()-Dinotefuran

890 (626–1264) 856 (590–1241) 1890 (1230–2890)

36.1 (24.9–52.2) 9.58 (6.79–13.52) 69.8 (47.1–103.4)

0.351 0.123 6.70

0.173 (0.104–0.287) 0.0545 (0.0396–0.0792) 2.67 (1.55–4.69)

a 95% confidence limits in parentheses. BGT, bungarotoxin; EPI, epibatidine. Reproduced with permission from Mori, K., Okumoto, T., Kawahara, N., Ozoe, Y., 2001. Interaction of dinotefuran and its analogues with nicotinic acetylcholine receptors of cockroach nerve cords. Pest Mgt Sci. 58, 190–196.

p0145

64 Neonicotinoid Insecticides

1. The tetrahydro-fur-2-ylmethyl derivative (shift of oxygen position) reflected diminished activity. 2. The tetrahydro-fur-3-yl moiety is important; the cyclopentanylmethyl (deoxy derivative), the 3-methoxy-N-propyl (open-ring ether) and the N-methyl-pyrrolidine-3-ylmethyl derivatives showed very low or negligible activity. 3. Introduction of a methyl group at the 4- or 5-position of the tetrahydro-fur-3-yl ring gave intermediate activity (Wakita et al., 2003), but introduction of methyl at the 2- or 3-position of the THF ring system and in a-position of the side chain decreases activity. An ethyl group reduced greatly activity even if it was at the 4-position (Wakita et al., 2003). The structural factors in the N-nitro guanidine moiety of (
)-dinotefuran were described as follows: 1. Deletion or extension to the alkyl group, like ethyl, of the terminal N-methyl group and addition of one more N-methyl group usually decrease the activity. 2. Carbocyclic cyclization to 5- and 6-ring systems maintains the activity. 3. Perhydro-1,3,5-oxadiazine 6-ring cyclization similar to that in thiamethoxam is inactive. 4. Modification of the N-nitroimino chromophore of dinotefuran to the nitromethylene, but not to the N-cyanoimino group maintains the insecticidal activity. These results indicate that the modification of this moiety of (
)-dinotefuran results in drastic changes in potency and that the incorporation of structural fragments known from previous neonicotinoid insecticides does not necessarily lead to compounds retaining higher activity (Mori et al., 2001). Also in this structural type the distance of three methylene chains between the nitrogen on the imidazolidine ring and the hydrogen acceptor oxygen was important (Wakita et al., 2003). 5.3.3.3. Bioisosteric Segments of Neonicotinoids

Retrospective considerations regarding bioisosteric segments of the developed and commercial neonicotinoids gave insight into general structural requirements (segments i–iii, Figure 4) for all the different ring systems and noncyclic structures (Jeschke et al., 2002). Several common molecular features, when comparing compounds with ring systems, like imidazolidine (imidacloprid), and its isosteric alternatives, like thiazolidine (thiacloprid), perhydro-1,3,5-oxadiazine (thiamethoxam), or hexahydro-1,3,5-triazine (Agro Kanesho’s AKD-1022) and the functional groups like N-nitroimino [›N NO2], N-cyanoimino w

[›N CN], or nitromethylene [›CH NO2] of these w w insecticides, have been described (Tomizawa et al., 2000). After superposition of the most active derivatives, it was possible to state the molecular shape similarity of the second generation neonicotinoids. It was found that electrostatic similarity of the most active compounds correlates well with the binding affinity (Nakayama and Sukekawa, 1998; Sukekawa and Nakayama, 1999), and a similar correlation was obtained by CoMFA (Nakayama, 1998; Okazawa et al., 1998). 5.3.3.3.1. Ring systems versus noncyclic structures In comparison to the corresponding ring systems, the noncyclic structures exhibit similar broad insecticidal activity by forming a so-called quasi-cyclic conformation when binding to the insect nAChR (Kagabu, 2003a). Thus, the three commercial noncyclic structures – nitenpyram, acetamiprid, and clothianidin – can be regarded as examples, if retrosynthetic considerations are carried out (Jeschke et al., 2002; Kagabu, 2003a). The noncyclic neonicotinoides are generally less lipophilic than the corresponding neonicotinoids with a ring structure (see Section 5.3.3.4). Based on the CoMFA results, a binding model for imidacloprid was described. This model clarified that the nitrogen of the CPM moiety interacts with a hydrogen-donating site of the nAChR, and that the nitrogen atom at the 1-position of the imidazolidine ring interacts with the negatively charged domain (Nakayama, 1998; Okazawa et al., 1998). Furthermore, the binding activity of noncyclic structures (e.g., acetamiprid, nitenpyram, and related compounds) to the nAChR of houseflies was measured and the results were analyzed using CoMFA. Superposition of stable conformations of nitenpyram, acetamiprid, and imidacloprid showed that the preferred regions for negative electrostatic potentials near the oxygen atoms of the nitro group, as well as the sterically forbidden regions beyond the imidazolidine 3-nitrogen atom of imidacloprid, were important for binding (Okazawa et al., 2000). The area around the 6-chloro atom of the CPM moiety was described as a sterically permissible region. Apparently the steric interactions were more important for noncyclic neonicotinoids than for the cyclic derivative, imidacloprid. Finally, it was also demonstrated that the noncyclic structures bind to the nAChR recognition site in a manner similar to ring structures like imidacloprid, and that the electrostatic properties of the noncyclic amino and cyclic imidazolidine structures affected their binding affinity (Figure 9). Figure 10 shows atom-based aligments of imidacloprid, clothianidin, dinotefuran and acetamiprid.

Neonicotinoid Insecticides

65

ring of dinotefuran differs dramatically from the other neonicotinoids under investigation. While the latter all show some large contributions at the aromatic moiety, the tetrahydrofuryl moiety seems to be much less attractive for nucleophilic attack.

Figure 9 Stable conformations and predicted properties of binding site: imidacloprid (white), nitenpyram (blue), acetamiprid (red). (Adapted from Akazawa et al., 2000.)

Figure 10 Alignment of DFT/BP/SVP/COSMO optimized geometries of imidacloprid, clothianidin, dinotefuran, and acetamiprid. The alignment was done by minimization of the mutual spatial distance of three pharmacophoric points, namely (i) the positively charged carbon atom connected to the ›N NO2 and w ›N CN moiety, respectively, (ii) the nitro/cyano groups themw selves, and (iii) the nitrogens of the aromatic rings and the oxygen of the tetrahydrofuran ring.

The aligment shown does not necessarily reflect the active conformation. However, the following arguments hold true for each of these family of conformers. The ion pairs of the aromatic nitrogen atoms in imidacloprid, clothianidin, and acetamiprid point in the same direction, and that this is also true for one of the two ion pairs of oxygen in dinotefuran. The tetrahydrofuryl ring of dinotefuran is more or less perpendicular to the heteroaromatic ring systems of the other neonicotinoids. Visual inspection of the nucleophilic Fukui functions (Figure 11) shows that the tetrahydrofuryl

5.3.3.3.2. Isosteric alternatives to the heterocyclic N-substituents The nitrogen-containing hetarylmethyl group as N-substituent (CPM, CTM) has a remarkably strong influence on the insecticidal activity. X-ray crystal structure analysis of imidacloprid and related neonicotinoids indicated that distances between the van der Waals surface of the CPM nitrogen and the atomic center of the pharmacophoric nitrogen are 5.45–6.06 A˚ (Tomizawa et al., 2000). This range coincides with the distance between the ammonium nitrogen and carbonyl oxygen of acetylcholine, and between the nitrogen atoms of (S)-nicotine (Kagabu, 1997a). Alternatively, the CPM and CTM moieties were assumed to be able to participate in hydrogen bonding, like the pyridine ring of (S)-nicotine, and that this is important for the insecticidal activity. The CTM substituent generally confers higher potency in the clothianidin and N-desmethyl-thiamethoxam series than the CPM moiety in the imidacloprid, thiacloprid, acetamiprid, and nitenpyram series (Zhang et al., 2000). Surprisingly, replacing both CPM and CTM by an oxygencontaining five-membered heterocycle resulted in a novel N-substituent TFM, that led to the development of the insecticide (
)-dinotefuran. It was found that the TFM structure can be taken as an isoster of the CPM and CTM moiety (Kagabu et al., 2002). In an attempt to understand this, the hydrogen bonding regions of CPM, CTM, and TFM were projected onto their respective Connolly surfaces (Jeschke et al., 2002) (Figure 12). 5.3.3.3.3. Bioisosteric pharmacophors The particularly high potency of the neonicotinoids bearing N-nitroimino, N-cyanoimino, or nitromethylene moieties, which have a negative electrostatic potential, implies a positive electrostatic potential for the corresponding insect nAChR recognition site (Nakayama and Sukekawa, 1998). Therefore, considerable attention has been given to the possible involvement of the pharmacophoric nitrogen in neonicotinoid action. In order to understand better the structural requirements, binding activity was analyzed using CoMFA (Akamatsu et al., 1997). SAR analyses have also been performed for in vitro activities (Nishimura et al., 1994). In particular, 3D QSAR procedures are helpful to predict the receptor–ligand interaction (Nakayama, 1998; Okazawa et al., 1998,

66 Neonicotinoid Insecticides

Figure 11 Isosurfaces for Fukui functions for nucleophilic attack of (a) imidacloprid, (b) clothianidin, (c) dinotefuran, and (d) acetamiprid. Three levels of isosurfaces are displayed: 0.005 (green, opaque), 0.001 (yellow, transparent), and 0.0005 (white, transparent).

Figure 12 Isosteric alternatives to the heterocyclic N-substituents. Connolly surfaces of the CPM moiety from imidacloprid, nitenpyram, acetamiprid (blue), CTM moiety from thiamethoxam and clothianidin (magenta), and TFM from (
)-dinotefuran (green). Atomic charges have been driven via the Mulliken partitioning scheme DFT/BP/TZVP/COSMO Kohn–Sham orbitals. (a) H-bonding potentials is mapped onto the Connolly surface; (b) atomic charges are projected onto the Connolly surface. (Reproduced with permission from Jeschke, P., Schindler, M., Beck, M., 2002. Neonicotinoid insecticides: retrospective consideration and prospects. Proc. BCPC: Pests and Diseases 1, 137–144.)

2000). As described in an alternative binding model for imidacloprid, the interatomic distance of 5.9 A˚ between the oxygen of the nitro group (at the van der Waals surface) and the nitrogen in 1-position was also noted as adequate (Kagabu, 1997a). That means the oxygen of the nitro group and the cyano nitrogen are well suited as acceptors for hydrogen bonding with the nAChR, in place of ring nitrogen

atoms in CPM and CTM or the ring oxygen in TFM. Thus the p-conjugated system composed of a N-nitroimino or N-cyanoimino group and the conjugated nitrogen in 1-position are considered essential moieties for the binding of neonicotinoids to the putative cationic subsite in insect nAChR (Figure 13). On the other hand, Maienfisch et al. (1997) reported that a nitroenamine isomer (N3-free

Neonicotinoid Insecticides

67

(Figure 14, Table 3). Due to the excellent systemic properties of neonicotinoids conferred by the moderate water solubility, these insecticides are effective against their main target pests, which are sucking insects, like aphids, leafhoppers, and whiteflies.

Figure 13 Binding to putative cationic subsite in insect nAChR (Kagabu, 1997a).

imidacloprid) posseses similar insecticidal activity to imidacloprid (Boe¨ lle et al., 1998). 5.3.3.4. Physicochemistry of Neonicotinoids

The physicochemical properties of the ring system and noncyclic neonicotinoids played an important role in their successful development. In addition, photostability is a significant factor in field performance of this class of insecticides (see Section 5.3.2). As described (Kagabu and Medej, 1995; Kagabu, 1997a; Kagabu and Akagi, 1997), the energy gap for the different functional groups [›X Y], from w the ground state to the single state, excited in the order [›N CN] > [›N NO2] > [›CH NO2]. w w w For practical application of neonicotinoids, such as for soil and seed treatment, as well as for foliar application, the uptake and translocation in plants is crucial for their insecticidal activity. Thereby, not only the bioisosteric segments of neonicotinoids (see Section 5.3.3.3.3) but also the whole molecular shape (Figure 12), including the resultant water solubility, has to be considered. Neonicotinoid insecticides are push–pull olefins made up of conjugated electron donating and accepting groups (Kagabu, 1997a). Such polar, nonvolatile molecules have a high water solubility (Table 3) and low POW values compared with other nonpolar insecticidal classes. The following characteristics were described from Kagabu (1997a): 1. Noncyclic neonicotinoids are less lipophilic than the corresponding ring system neonicotinoids. 2. Concerning the functional group [›X Y] as w part of the pharmacophore [ N C(E)›X Y] w w w (iii), water solubility increases in the order of [›CH NO2] > [›N CN] > [›N NO2]. w w w 3. Regarding E, the lipophilicity increased in the order of S > C > O > NH (Kagabu, 1996). Somewhat more lipophilic neonicotinoids should be better for seed treatment application, because uptake by roots is more effective than in the case of more hydrophilic compounds (Briggs et al., 1982)

5.3.3.4.1. Physicochemical properties of commercialized neonicotinoids The physicochemical properties of a compound are related to its structural formula. However, it is not possible to predict the behavior of a substance, with sufficient certainity, only on the basis of its formula and physicochemical properties (Stupp and Fahl, 2003). 5.3.3.4.1.1. Imidacloprid Due to the special moieties like the CPM residue and the 2-(N-nitroimino)imidazolidine 5-ring system, imidacloprid has very weak basic properties under environmental conditions (see Section 5.3.8.1.2). Water solubility and low partition coefficient in octanol–water are not influenced by pH values between 4 and 9, and at 20  C (Krohn and Hellpointer, 2002) (Table 3). The low partition coefficient of imidacloprid indicates that it has no potential to accumulate in biological tissues and further enrichment in the food chain. The occurrence of imidacloprid in the air is determined by its low vapor pressure of 4  1010 Pa which excludes volatilization from treated surfaces. The rapid uptake and translaminar transport of imidacloprid from the treated upper leaf side to the lower surface is excellent, as observed in cabbage leaves (Elbert et al., 1991), and in rice and cucumber (Ishii et al., 1994). However, imidacloprid has a considerable acropetal mobility in xylem of plants. In contrast, its penetration and translocation in cotton leaves was less pronounced, as qualitatively reflected by phosphor-imager autoradiography (Buchholz and Nauen, 2001). This xylem mobility makes imidacloprid especially useful for seed treatment and soil application, but it is equally effective for foliar application (Elbert et al., 1991). Due to its lack of any acidic hydrogen, the pKa of imidacloprid is >14, and therefore its transport within the phloem is unlikely, as been shown in several studies (Stein-Do¨ necke et al., 1992; Tro¨ ltzsch et al., 1994). Systemic properties were examined using 14C-labeled imidacloprid. The translocation of imidacloprid in winter wheat and its uptake and translocation from treated cotton seeds into the growing parts of the cotton plant is described by Elbert et al. (1998). Registered patterns of use of imidacloprid in agriculture now include traditional foliar spray application as well as soil drench application, drip irrigation, trunk (injection) application, stem or granular treatments and seed treatment.

Table 3 Physicochemical properties of commercialized neonicotinoids Ring systems

Noncyclic structures

Physical and chemical properties

Imidacloprid

Thiacloprid

Thiamethoxam

Nitenpyram

Acetamiprid

Clothianidin

Color and physical state

Colorless crystals

Slightly cream/ crystalline powder

Pale yellow crystals

Colorless crystals

Clear/coloress solid powder

Melting point ( C) Henry’s law constant (Pa  m3 mol1) (at 20  C) Density (g ml1) (at 20  C) Vapor pressure (Pa) (at 25  C) (at 20  C) Solubility in water (g l1) (at 20  C) Solubility in organic solvents (g l1 at 25  C) Dichloromethane N-hexane N-heptane Methanol Ethanol Xylene Toluene 1-Octanol Acetone Acetonitrile Ethyl acetate Partition coefficient in octanol– water (at 25  C) log Pow

144 2  1010

Yellow/ crystalline powder 136

139.1

83–84

98.9 2000 (m/f)

1563 >2000

1680 (m) 1575 (f) >2000 (m/f)

217 (m) 146 (f)

>5000 (m/f) >2000 (m/f)

2804 (m) 2000 (f)

Rat

>5323 (dust)

3720

>5800 (m/f)

>0.29 (m/f)

>6141 (m/f)

Rabbit

No irritation

>2535 (m) 1223 (f) No irritation

No irritation

No irritation

No irritation

No irritation

Eye irritation

Rabbit

None

No irritation

No irritation

Very slight

No irritation

No irritation

Skin sensitation

Guinea pig

No skin sensitation

No skin sensitation

No skin sensitation

No skin sensitation

a

Elbert et al. (1990). Elbert et al. (2000). c Maienfisch et al. (1999a). d Tomlin (2000). e Ohkawara et al. (2002). m, males; f, females; a.i., active ingredient; bw, body weight. b

No skin sensitation

Acute percutaneous Acute percutaneous No skin sensitation

t0035

Table 7 Environmental profile of commercialized neonicotinoids (selected relevant data) Ring systems Test species (acute toxicity test)

Noncyclic structures

Imidacloprid a

Thiacloprid b

Thiamethoxamc

Nitenpyramd,g

>5000 (food) 152 31

>5000 (food)g 2716 49

>5200 (food) 1552

>5620 (food) 2250 (capsule)

211 (96 h)

29.6 (96 h) 24.5 (96 h)

>125 (96 h) >114 (96 h)

Acetamiprid e

Clothianidinf

(
)-Dinotefurane

>5200 (food) >2000

1000

Birds

Mallard duck (LC50 mg a.i. kg1 diet) Bobwhite quail (LD50 mg a.i. kg1 bw) Japanese quail (LD50 mg a.i. kg1 bw)

180

>2000

Fish

Rainbow trout (LC50 mg a.i. l1) Bluegill sunfish (LC50 mg a.i. l1) Carp (LC50 mg a.i. l1) Invertebrates Water flea, Daphnia magna

280 (96 h)

>10 (96 h)

>100 (96 h) >120 (96 h)

>1000 (96 h)

>100

>1000 (96 h)

85 (48 h)

>85.1 (48 h)

>100 (48 h)

>100 (48 h)h

>1000

>120 (48 h)

10.7 (14 days)

105 (14 days)g

>1000 (14 days) (soil)

32.2 (14 days)

>1000

13.21 (14 days) (dry soil)

1

(EC50 mg a.i. l ) Earthworm, Eisenia foetida (LC50 mg a.i. kg1 dry soil) a

Pflu¨ger and Schmuck (1991). Schmuck (2001). c Maienfisch et al. (1999a). d Akayama and Minamida (1999). e Tomlin (2000). f Ohkawara et al. (2002). g Technical material. h Unpublished data; Sumitomo Chemical Takeda Agro, 1999. b

>40 (48 h)

1000 (48 h)

86 Neonicotinoid Insecticides

p0520

occurrence of imidacloprid in the air is determined by its low vapor pressure of 4  1010 Pa, which excludes volatilization from treated surfaces (Krohn and Hellpointer, 2002). Thiacloprid possesses a favorable environmental profile and will disappear from the environment after having been applied as a crop protection chemical. The effect of technical and formulated (SC 480) thiacloprid on respiratory (carbon turnover) and nitrogen mineralization rates in soil were examined in a series of 28-day studies on soil microflora (Schmuck, 2001). Thiacloprid had no adverse effect either on the microbial mineralization of nitrogen in different soil types like silty sand (0.6% org.C, pHKCl 5.7) or loamy silt soil (2.3% org.C, pHKCl 7.1) after addition of lucerne–grass–green meal (5 g kg1) and at both the maximum and 10  application rates (Schmuck, 2001). The half-lives in soil measured under field conditions of northern Europe ranged from 9 to 27 days, and in southern Europe from 10 to 16 days (Krohn, 2001). The major metabolites formed are only intermediates before complete mineralization to carbon dioxide occurs. Under practical field conditions thiacloprid is not toxic to earthworms (Eisenia foetida) or soil microorganisms (Table 7). Furthermore, thiacloprid and its formulations pose a favorably low toxicity hazard to honeybees and bumblebees. Consequently, the compound can be applied to flowering crops at the recommended rates without posing a risk to bees (CalypsoTM up to 200 g a.i. ha1). In addition, CalypsoTM does not affect the pollination efficacy of pollinators (Elbert et al., 2000). Thiacloprid must be classified as being only slightly mobile in soil, and hence it has low potential for leaching into groundwater. In the aquatic environment thiacloprid will also undergo rapid biotic degradation, with a half-life of 12–20 days (Krohn, 2001). Because of its low vapor pressure (3  1010 Pa) and its water solubility of 0.185 g l1, the potential for its volatilization is negligible. Acute and chronic toxicity testing in fish and water-fleas (Daphnia magna) indicates a very low to negligible risk to these species following accidental contamination with thiacloprid by spray drift. Reproduction of water-fleas and fish was also not affected at environmentally relevant water concentrations of thiacloprid. Other aquatic invertebrates revealed a higher sensitivity to thiacloprid, especially insect larvae and amphipod species such as Chironomus and Hyallela. Insect larvae were also identified as the most sensitive species in a pond study. However, the rapid dissipation of thiacloprid in water bodies allows for a rapid recovery of affected aquatic insect populations. Algae, represented by

Scenedesmus subspicatus and Pseudokirchneriella subcapitata, are not sensitive to thiacloprid. Toxicity exposure ratio values indicate a high margin of safety for algal species. Harmful concentrations of thiacloprid or its metabolites are most unlikely in aquatic ecosystems. There is no potential for bioconcentration in fish due to the log POW of 1.26 for thiacloprid. The metabolites occurring in groundwater are of no concern as noted by toxicological, ecotoxicological, and biological testing. In laboratory soils, thiamethoxam degrades at moderately slow rates (Maienfisch et al., 1999a). The compound is photolyzed rapidly in water. In natural aquatic systems, e.g., rice paddies, degradation also occurs in the absence of light by microbial degradation. Based on the low vapor pressure (Table 3) and the results of soil volatility studies, significant volatilization is not expected. Thiamethoxam has a low toxicity by ingestion to birds, and is practically nontoxic to fish, Daphnia and molluscs. In addition, the earthworm species E. foetida and green algae were also found to be insensitive (Maienfisch et al., 2001a). Thiamethoxam can be classified as slightly to moderately harmful to most beneficial insects, but safe to predatory mites in the field (Maienfisch et al., 1999a). Alternatively, thiamethoxam and all other nitroguanidines have to be considered toxic to bees. However, thiamethoxam showed no bioaccumulation potential, and is moderately mobile in soil and degrades at fast to moderate rates under field conditions (Maienfisch et al., 1999a). Nitenpyram rapidly decomposed in water, under the conditions of 12 000 lux irradiation intensity using a 500 W xenon lamp (half-life at pH 5, 18–21 min; in purified water, 16 min), and in soil under aerobic conditions (half-lives flooded 100 mg l1 and >1000 mg l1, respectively (Yamada et al., 1999). Acetamiprid shows less adverse effects on both natural and enemies, such as predaceous mites, and beneficial insects, as well as honeybee and bumblebee (Takahashi et al., 1998). Chlothianidin degraded moderately under field conditions and was found to be a neonicotinoid of medium mobility based on laboratory experiments (Stupp and Fahl, 2003). With regard to its behavior in aquatic environments, photolysis contributes significantly to degradation, and clothianidin finally mineralizes to carbondioxide. Its degradation in water/sediment systems under aerobic conditions is moderate. The main part of the applied active

p0525

p9000

Neonicotinoid Insecticides

substance is bound irreversibly to the sediment, thus avoiding subsequent contamination (Stupp and Fahl, 2003). In addition, in a water/sediment study degradation of clothianidin was observed to be significantly faster (factor 2–3) under anaerobic conditions than in aerobic conditions (Stupp and Fahl, 2003). Based on the physicochemical properties of clothianidin (see Section 5.3.3.4) no volatilization, and thus, no significant amounts of this compound can be expected in the atmosphere. Therefore clothianidin does not appear to represent a risk for the environment.

5.3.9. Resistance Resistance in arthropod pest species comprises a change in the genetic composition of a population in response to selection by pesticides, such that control of the pest species in the field may be impaired at the recommended application rates. Selection for resistance in pest populations (including weeds, fungi, mites, and insects) due to frequent applications of agrochemicals has been one of the major problems in modern agriculture. Resistance to insecticides and acaricides appeared very early, and the extent and economic impact remain greater than for other agrochemicals. Between the beginning of the last century and the mid-1950s, the number of resistant arthropod species grew gradually with only a few resistant species described per decade. However, this rate increased markedly after the introduction of the organophosphates to more than 30 new resistant species every 2 years through the early 1980s. Ten years later, more than 500 arthropod species were known to be resistant to at least one insecticide or acaricide (Georghiou, 1983; Green et al., 1990). Among them are 46 hopper and aphid species, which show in some cases very high levels of resistance to conventional types of insecticide chemistry, i.e., organophosphates, and carbamate and pyrethroid insecticides. The invention and subsequent commercial development of neonicotinoid insecticides, such as imidacloprid, has provided agricultural producers with invaluable new tools for managing some of the world’s most destructive crop pests, primarily those of the order Hemiptera (aphids, whiteflies, and planthoppers) and Coleoptera (beetles), including species with a long history of resistance to earlier-used products (Figure 20). However, the speed and scale with which imidacloprid, the commercial forerunner of neonicotinoids, was incorporated into control strategies around the world prompted widespread concern over the development of imidacloprid resistance (Cahill et al., 1996; Denholm et al., 2002).

87

To a large extent these pessimistic forecasts have not been borne out in practice. Imidacloprid has proved remarkably resilient to resistance, and cases of resistance that have been reported are still relatively manageable and/or geographically localized. The existence of strong resistance in some species has nonetheless demonstrated the potential of pests to adapt and resist field applications of neonicotinoids. The ongoing introduction of new molecules (e.g., acetamiprid, thiamethoxam, nitenpyram, thiacloprid, and clothianidin), unless carefully regulated and coordinated, seems bound to increase exposure to neonicotinoids, and to enhance conditions favoring resistant phenotypes. 5.3.9.1. Activity on Resistant Insect Species

The incidence and management of insect resistance to neonicotinoid insecticides was recently reviewed by Denholm et al. (2002). Resistance to neonicotinoid insecticides is still rare under field conditions and baseline susceptibility data have been provided in the past and more recently especially for imidacloprid in order to monitor for early signs of resistance in some of the most destructive pest insects (Cahill et al., 1996; Elbert et al., 1996; Elbert and Nauen, 1996; Foster et al., 2003; Nauen and Elbert, 2003; Rauch and Nauen, 2003; Weichel and Nauen, 2003). Baseline susceptibility data and derived diagnostic concentrations to monitor resistance are usually calculated from composite log-dose probitmortality lines, including the combined curves of several strains of a certain pest collected from all over the world or at least different parts of the world where the compound is supposed to be used. A compilation for imidacloprid of such data is given for some of the most important pests, i.e., Myzus persicae, Aphis gossypii, Phorodon humuli, Bemisia tabaci, Trialeurodes vaporariorum, and Leptinotarsa decemlineata (Table 8). There have been only low levels of tolerance detected in European and Japanese samples of M. persicae (Devine et al., 1996; Nauen et al., 1996), but it was not possible to link this tolerance to specific biochemical markers (Nauen et al., 1998a). In most cases the lower susceptibility to imidacloprid and other neonicotinoids was correlated with a decreased efficacy of nicotine (Devine et al., 1996; Nauen et al., 1996). Furthermore differences in hardiness were sometimes observed in field strains bioassayed directly upon receipt, which allow strains to survive longer in comparison to susceptible laboratory populations, when exposed to imidaclopridtreated leaves (Nauen and Elbert, 1997). However, such effects disappeared when the exposure time (especially in systemic bioassays) was extended

88 Neonicotinoid Insecticides

Table 8 Baseline susceptibility of some high risk pests to imidacloprid Species

Diagnostic dose

Bioassay system/assessment time

Reference

Myzus persicae Myzus persicae Aphis gossypii Phorodon humuli Bemisia tabaci Bemisia tabaci Leptinotarsa decemlineata Leptinotarsa decemlineata

15 ppm 2.25 ng per aphid 13 ppm 13 ppm 16 ppm 1 ppm 8 ppm 0.2 mg per beetle

Leaf dip (6-well plate) Topical Leaf dip (6-well plate) Leaf dip (6-well plate) Systemic bioassay Leaf dip Artificial diet (larvae) Topical (adult)

Nauen and Elbert (2003) Foster et al. (2003) Nauen and Elbert (2003) Weichel and Nauen (2003) Cahill et al. (1996) Rauch and Nauen (2003) Olson et al. (2000) Nauen (unpublished data)

from 48 h to 72 h, and also after maintaining such strains under laboratory conditions for some weeks (Nauen and Elbert, 1997). More recently Foster et al. (2003b) demonstrated that tolerance to imidacloprid in M. persicae from different regions in Europe also provided cross-tolerance to acetamiprid. The authors were able to show a clear correlation between ED50 values of acetamiprid and imidacloprid for strains with a different degree of tolerance. However, tolerance factors compared to a susceptible reference population never exceeded factors of 20, and field failures were not seen (Foster et al., 2003b). One species of major concern over the last decade is the tobacco or cotton whitefly, B. tabaci; this is a serious pest in many cropping systems worldwide and several biotypes of this species have been described (Perring, 2001). The most widespread biotype is the B-type, which is also known as B. argentifolii Bellows & Perring. The B-type whitefly is a common pest, particularly in cotton, vegetables, and ornamental crops, both by direct feeding and as a vector of numerous plant pathogenic viruses. In southern Europe, it coexists with another biotype, the Q-type, which was originally thought to be restricted to the Iberian peninsula, but which is now also known to occur in some other countries throughout the Mediterranean area, including Italy and Israel (Brown et al., 2000; Palumbo et al., 2001; Nauen et al., 2002; Horowitz et al., 2003). The biotypes B and Q can easily be distinguished by randomly amplified polymorphic DNA polymerase chain reaction (RAPD-PCR) or native polyacrylamide gel electrophoresis (PAGE) and subsequent visualization of their nonspecific esterase banding pattern (Guirao et al., 1997; Nauen and Elbert, 2000). As a consequence of extensive exposure to insecticides, B. tabaci has developed resistance to a wide range of chemical control agents (Cahill et al., 1996). The need for a greater diversity of chemicals for whitefly control in resistance management programs has been met by the introduction of several insecticides with new modes of action, which are

unaffected by mechanisms of resistance to organophosphates or pyrethroids. Since the introduction of imidacloprid, the neonicotinoids have been the fastest-growing class of insecticides. Imidacloprid exhibits an excellent contact and systemic activity and therefore has been largely responsible for the sustained management of B. tabaci in horticultural and agronomic production systems worldwide. Beside imidacloprid, there are other neonicotinoids with good efficacy against whiteflies, e.g., acetamiprid and thiamethoxam. In Israel, monitoring of resistance in B. tabaci to imidacloprid and acetamiprid was initiated in 1996 in cotton and greenhouse ornamental crops. After 2 years of use in cotton, no apparent resistance to imidacloprid and acetamiprid was reported (Horowitz et al., 1998). However, 3 years of acetamiprid use in greenhouses in Israel resulted in a 5–10-fold decrease in susceptibility of B. tabaci to acetamiprid (Horowitz et al., 1999). In the past only a few cases of lowered neonicotinoid susceptibility in B-type B. tabaci have been described, among them strains from Egypt and Guatemala that were recently reported (El Kady and Devine, 2003; Byrne et al., 2003). In Arizona, where imidacloprid has been used since 1993, monitoring of B. tabaci populations from cotton fields, melon fields, and greenhouse vegetables suggested reduced susceptibility to imidacloprid from 1995 to 1998, but subsequent monitoring showed that these populations had actually regained and sustained susceptibility to imidacloprid in 1999 and 2000 (Li et al., 2000, 2001). Furthermore, imidacloprid use in Arizona and California remains high, but no signs of reduced control in the field have been reported yet (Palumbo et al., 2001). B-type whiteflies have been shown to develop resistance to imidacloprid under selection pressure in the laboratory (Prabhaker et al., 1997). There was a moderate increase of resistance of up to 17-fold in the first 15 generations, but 82-fold resistance after 27 generations. However, resistance was not stable and disappeared after a few generations without insecticide pressure. Resistance to

Neonicotinoid Insecticides

imidacloprid conferring a high level of crossresistance to thiamethoxam and acetamiprid was first demonstrated, and best studied in Q-type B. tabaci from greenhouses in the Almeria region of southern Spain, but was also detected in single populations from Italy and recently Germany as well (Nauen and Elbert, 2000; Nauen et al., 2002; Rauch and Nauen, 2003). Neonicotinoid resistance seem to remain stable in all field-collected Q-type strains maintained in the laboratory without further selection pressure (Nauen et al., 2002). Neonicotinoid cross-resistance was also reported in B-type whiteflies from cotton in Arizona but at lower levels (Li et al., 2000). More recently a high level of cross-resistance between neonicotinoids was also described in a B-type strain of B. tabaci from Israel, and resistance factors detected in a leaf-dip bioassay exceeded 1000-fold (Rauch and Nauen, 2003). The Colorado potato beetle, L. decemlineata has a history of developing resistance to virtually all insecticides used for its control. The first neonicotinoid, i.e., imidacloprid was introduced for controlling Colorado potato beetles in North America in 1995. Concerns over resistance development were reinforced when extensive monitoring of populations from North America showed about a 30-fold variation in LC50 values from ingestion and contact bioassays against neonates (Olsen et al., 2000). Much of this variation appeared unconnected with imidacloprid use, and was probably a consequence of cross-resistance from chemicals used earlier. Lowest levels of susceptibility occurred in populations from Long Island, New York, an area that has experienced the most severe resistance problems of all with L. decemlineata. Zhao et al. (2000) and Hollingworth et al. (2002) independently studied single strains collected from different areas in Long Island, both treated intensively with imidacloprid between 1995 and 1997. In the first study, grower’s observations of reduced control were supported by resistance ratios for imidacloprid of 100-fold and 13-fold in adults and larvae, respectively. The second study reported 150-fold resistance from topical application bioassays against adults. In this case the strain was also tested with thiamethoxam, which had not been used for beetle control at the time of collection. Interestingly, resistance to thiamethoxam (about threefold) was far lower than to imidacloprid. Other reports referring to resistance to neonicotinoid insecticides were on species of lesser importance, including species either from field-collected populations or artificially selected strains. Among these were the small brown planthopper, Laodelphax striatellus (Sone et al., 1997), western flower

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thrips, Franklienella occidentalis (Zhao et al., 1995), houseflies, Musca domestica and German cockroach, Blattella germanica (Wen and Scott, 1997), Drosophila melanogaster (Daborn et al., 2001), Lygus hesperus (Dennehy and Russell, 1996), and brown planthoppers, Nilaparvata lugens (Zewen et al., 2003). 5.3.9.2. Mechanisms of Resistance

Many pest insects and spider mites have developed resistance to a broad variety of chemical classes of insecticides and acaricides, respectively (Knowles, 1997; Soderlund, 1997). One of the three major classes of mechanisms of resistance to insecticides in insects is allelic variation in the expression of target proteins with modified insecticide binding sites, e.g., acetylcholinesterase insensitivity towards organophosphates and carbamates, voltage-gated sodium channel mutations responsible for knockdown resistance to pyrethroids, and a serine to alanine point mutation (rdl gene) in the g-aminobutyric acid (GABA)-gated chloride channel (GABAA-R) at the endosulfan/fipronil/dieldrin binding site (ffrenchConstant et al., 1993; Williamson et al., 1993, 1996; Mutero et al., 1994; Feyereisen, 1995; Dong, 1997; Soderlund, 1997; Zhu and Clark, 1997; Bloomquist, 2001; Gunning and Moores, 2001; Siegfried and Scharf, 2001). The second – and often most important – class of resistance mechanisms in insect pest species is metabolic degradation involving detoxification enzymes such as microsomal cytochrome P-450 dependentmonooxygenases, esterases, andglutathione S-transferases (Hodgson, 1983; Armstrong, 1991; Hemingway and Karunarantne, 1998; Berge´ et al., 1999; Devonshire et al., 1999; Feyereisen, 1999; Hemingway, 2000; Field et al., 2001; Scott, 2001; Siegfried and Scharf, 2001). The third, least important mechanism is an altered composition of cuticular waxes which affects penetration of toxicants. Reduced penetration of insecticides through the insect cuticle has often been described as a contributing factor, in combination with target site insensitivity or metabolic detoxification (or both), rather than functioning as a major mechanism on its own (Oppenoorth, 1985). Most of the mechanisms mentioned above affect in many cases the efficacy of more than one class of insecticides, i.e., constant selection pressure to one chemical class could to a greater or lesser extent confer crossresistance to compounds from other chemical classes (Oppenoorth, 1985; Soderlund, 1997). When the first neonicotinoid insecticide was introduced to the market in 1991, aphids were considered to be high risk pests with regard to their potential to develop resistance to this class of

90 Neonicotinoid Insecticides

chemicals. They have a high reproductive potential, and extremely short life cycle allowing for numerous generations in a growing season. Combined with frequent applications of insecticides that are usually required to maintain aphid populations below economic thresholds, resistance development is facilitated in these species, resulting in control failures. Such control failures have been reported for organophosphorus compounds for many decades, and more recently also for pyrethroids (Foster et al., 1998, 2000; Devonshire et al., 1999; Foster and Devonshire, 1999). One of the major aphid pests is the green peach aphid, M. persicae. Resistance of M. persicae to insecticides is conferred by increased production of a carboxylesterase, named E4 or FE4, which provides cross-resistance to carbamates, organophosphorus, and pyrethroid insecticides (Devonshire and Moores, 1982; Devonshire, 1989). This esterase overproduction was shown to be due to gene amplification (Field et al., 1988, 2001). It was the sole resistance mechanism reported in M. persicae for more than 20 years, and only recently an insensitive (modified) acetylcholinesterase was described as a contributing factor in carbamate resistance in M. persicae (Moores et al., 1994a, 1994b; Nauen et al., 1996). The insensitive acetylcholinesterase in M. persicae confers strong resistance to pirimicarb, and a little less to triazamate (Moores et al., 1994a, 1994b; Buchholz and Nauen, 2001). Due to the improvement of molecular biological techniques, Martinez-Torres et al. (1998) recently showed that knockdown resistance to pyrethroids, caused by a point mutation in the voltage-gated sodium channel, is also present in M. persicae. In summary, resistance to all major classes of aphicides occurs in M. persicae; however, the only class of insecticides not yet affected by any of the mechanisms described above are the neonicotinoids, including its most prominent member imidacloprid (Elbert et al., 1996, 1998a; Nauen et al., 1998a; Horowitz and Denholm, 2001). The most comprehensive studies on the biochemical mechanisms of resistance to neonicotinoid insecticides using an agriculturally relevant pest species were performed in whiteflies, B. tabaci (Nauen and Elbert, 2000; Nauen et al., 2002; Rauch and Nauen, 2003; Byrne et al., 2003). Biochemical examinations revealed that neonicotinoid resistance in Q-type B. tabaci collected in 1999 was not associated with a lower affinity of imidacloprid to nAChRs in whitefly membrane preparations (Nauen et al., 2002). This was confirmed more recently by testing strains ESP-00, GER-01, and ISR-02 obtained in the years 2000–2002 by Rauch and Nauen

(2003). Although neonicotinoid resistance was very high in these strains (up to 1000-fold), the authors found just a 1.3-fold and 1.7-fold difference in binding affinity between strains, and concluded that target site resistance is not involved in neonicotinoid resistance in those strains investigated. Piperonyl butoxide, a monooxygenase inhibitor, is generally used as a synergist to suppress insecticide resistance conferred by microsomal monooxygenases. Experiments with whiteflies pre-exposed to piperonyl butoxide suggested a possible involvement of cytochrome P-450 dependent monooxygenases in neonicotinoid resistance (Nauen et al., 2002). Rauch and Nauen (2003) biochemically confirmed that whiteflies resistant to neonicotinoid insecticides showed a high microsomal 7-ethoxycoumarin O-deethylase activity, i.e., up to eightfold higher compared with neonicotinoid susceptible strains. Furthermore, this enhanced monooxygenase activity could be correlated with imidacloprid, thiamethoxam, and acetamiprid resistance. Significant differences between glutathione S-transferase and esterase levels were not found between neonicotinoid resistant and susceptible strains of B. tabaci (Rauch and Nauen, 2003). Several metabolic investigations in plants and vertebrates showed that imidacloprid and other neonicotinoids undergo oxidative degradation, which may lead to insecticidally toxic and nontoxic metabolites (Araki et al., 1994, Nauen et al., 1999b; Schulz-Jander and Casida, 2002). Metabolic studies in B. tabaci in vivo revealed that the main metabolite in neonicotinoid-resistant strains is 5-hydroxy-imidacloprid, whereas no metabolism could be detected in the susceptible strain (Figure 23) (see Section 5.3.7.1.1). One can therefore suggest that oxidative degradation is the main route of imidacloprid detoxification in neonicotinoid resistant Q-type whiteflies (Rauch and Nauen, 2003). Compared to imidacloprid, the 5-hydroxy metabolite showed a 13-fold lower binding affinity to whitefly nAChR. This result was in accordance with previous studies with head membrane preparations from the housefly (Nauen et al., 1998). The binding affinity expressed as the IC50 was highest with olefine (0.25 nM) > imidacloprid (0.79) > 5-hydroxy (5 nM) > 4-hydroxy (25 nM) > dihydroxy (630 nM) > guanidine and urea (>5000 nM). The biological efficacy in feeding bioassays with aphids correlated also with the relative affinities of the metabolites towards the housefly nAChR (Nauen et al., 1998b). The lower binding affinity of 5-hydroxy-imidacloprid compared to imidacloprid coincides with its lower efficacy against B. tabaci in the sachet test (17-fold). These data show that differences between binding to the

Neonicotinoid Insecticides

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nAChR are comparable to the efficacy differences in adult bioassays (Rauch and Nauen, 2003). Daborn et al. (2001) described D. melanogaster mutants exhibiting an eight- and sevenfold resistance to imidacloprid and DDT, respectively, which showed an overexpression of the P-450 gene Cyp6g1. However, it was not tested whether recombinant Cyp6g1 itself can metabolize imidacloprid; potentially it could metabolize a trans-acting factor which then upregulates presently uncharacterized P-450 genes. However, it was demonstrated that the recombinant cytochrome P-450 isozyme CYP3A4 from human liver can selectively metabolize the imidazolidine moiety of imidacloprid, resulting in 5-hydroxy-imidacloprid as the major metabolite (Schulz-Jander and Casida, 2002). It would therefore be fruitful to investigate if homologs of Cyp6g1 and Cyp3a4, or similar genes, are present in the B. tabaci genome, and if overexpression of these genes is associated with neonicotinoid resistance. Although considerable resistance to neonicotinoid insecticides was also reported for populations of the Colorado potato beetle from Long Island, NY, the underlying mechanisms associated have not been very well studied. Pharmacokinetic investigations on susceptible and resistant strains did not reveal any differences in the uptake and excretion of 14 C-labeled imidacloprid, nor were there any differences in metabolic conversion found (Hollingworth et al., 2002). The authors suggested a modification of the target site to be responsible for the observed resistance. But, preliminary investigations revealed no differences between resistant and susceptible strains in receptor binding studies using [3H]imidacloprid (Nauen and Hollingworth, unpublished data). Resistance development to imidacloprid would be disastrous in many regions, so there is a demand for an effective resistance management program, and interest in finding and developing new active ingredients for pest control. 5.3.9.3. Resistance Management

Monitoring and detection of insecticide resistance in order to recommend and implement effective resistance management strategies is currently one of the most important areas of applied entomology. This program is necessary to sustain the activity of as many active ingredients with different modes of action as possible by using alternate spray regimes, rotation, and sophisticated application techniques (Denholm and Rowland, 1992; McKenzie, 1996; Denholm et al., 1998, 1999; Horowitz and Denholm, 2001). Historically, the problem of

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insecticide resistance was tackled by continuously introducing new active ingredients for pest insect control. The number of insecticides available is high, but the biochemical mechanisms targeted by all these compounds is rather limited (Casida and Quistad, 1998; Nauen and Bretschneider, 2002). A pronounced decline in the introduction of new insecticides since the late 1970s revealed the limitations of a strategy relying on the continuous introduction of new active compounds (Soderlund, 1997). This problem has been recognized by the regulatory authorities; and the European Plant Protection Organization (EPPO) recently published guidelines outlining the background work on resistance required for registration of a new active ingredient (Heimbach et al., 2000). These requirements, which are provided by agrochemical companies as an essential part of the registration dossier, include . baseline susceptibility studies (testing several strains of a target species known to be prone to resistance development); . monitoring (continuous studies on the development of resistance of target species by simple bioassays after the launch of a new compound); and . possible resistance management strategies (how the new compound should be combined with others in order to expand its life time in the field).

5.3.10. Applications in Nonagricultural Fields Because of the high insecticidal efficacy together with its nonvolatility and stability under storage conditions, imidacloprid (PremiseÕ) has been successfully applied as a termiticide ( Jacobs et al., 1997a; Dryden et al., 1999), and has also been used for control of turf pests such as white grubs (Elbert et al., 1991). Furthermore, imidacloprid is the active ingredient of the insecticide MeritÕ, and is commonly incorporated into fertilizers for early control of grubs in turf (Armbrust and Peeler, 2002). In addition, imidacloprid was the first neonicotinoid to be used in a gel bait formulation for cockroach control. The imidacloprid gel showed outstanding activity even after 27 months’ under various conditions (Pospischil et al., 1999). As an endoparasiticide imidacloprid exerted activity against the gastrointestinal nematode Haemonchus contortus in sheep only at higher concentrations (Mencke and Jeschke, 2002). Synergistic mixtures containing imidacloprid are patented and these are useful against textile-damaging insects, such as

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92 Neonicotinoid Insecticides

moth (Tineola bisselliella, Tinea pellionella) and beetles (Attagenus, Anthrenus) (Mencke and Jeschke, 2002). Salmon-parasitizing crabs are controlled by addition of 100 ppm imidacloprid to seawater (Mencke and Jeschke, 2002). Due to its toxicological properties – favorable mammalian safety characteristics (Yamamoto et al., 1995) (Table 6), the absence of eye/skin irritation and skin sensitization potential – imidacloprid has been developed for control of lice in humans (Mencke and Jeschke, 2002) and veterinary medicine (Werner et al., 1995). Imidacloprid (worldwide trademark: AdvantageÕ) is the first neonicotinoid to have been developed for topical application in animals (Griffin et al., 1997) (see Section 5.3.10.1). Nitenpyram (CapstarÕ), a fast-acting, orally administered flea treatment, is absorbed into blood of the host animal, and is thus readily available for uptake by feeding fleas (Rust et al., 2003). Therefore, administration of nitenpyram is effective in eliminating adult fleas for up to 48 h after treatment. 5.3.10.1. Imidacloprid as a Veterinary Medicinal Product

The cat flea (Ctenocephalides felis), the primary ectoparasite of companion animals worldwide, will feed on a wide variety of animals in addition to cats and dogs (Rust and Dryden, 1997), although it is not equally well adapted to all hosts (Williams, 1993). Fleas threaten the health of humans and animals due to bite reactions and transmission of diseases (Kra¨ mer and Mencke, 2001), and in addition are major nuisance pests. Therefore, flea control, is necessary. In veterinary medicine, fleas are the primary cause for flea allergic dermatitis. This dermatitis results when the adult flea injects saliva into the host during blood feeding, which accelerates immunological response, leading to secondary infections of the skin. The last aspect of veterinary importance is the role of fleas in disease, for example, transmission of the cestode Dipylidium caninum. The role of fleas in human disease transmission has been known since historic times. The Oriental rat flea (Xenopsylla cheopis) is the major transmitter of Yersinia pestis, the bacterium that causes the bubonic plague in humans. The cat flea is also capable of transmitting Y. pestis, and human plague cases and even deaths associated with infected cats and dogs have been occasionally reported (Rust et al., 1971). Furthermore, a variety of bacteria and viruses have been reported to be transmitted by the dog flea (C. canis) as well as the cat flea (C. felis) (Kra¨ mer and Mencke, 2001). Moreover, cat owners have a high incidence of cat scratch fever, a zoonotic

disease caused by the Gram-negative bacterium Bartonella henselae. Recent research showed that flea feces are the major means of disease transmission between cats, and from cats to humans (Malgorzata et al., 2000). 5.3.10.1.1. Insecticidal efficacy in veterinary medicine Recommendations for the treatment of fleas on companion animals, and the selection of an insecticide and its formulation, are generally based upon the species and age of the animal to be treated, the level of infestation, the rate of potential reinfestation, and the thoroughness of environmental treatment. However, the selection of an insecticide formulation by a pet owner is actually based on economics and the product’s ease of use (Williams, 1993). Another factor in the choice of a flea treatment by pet owners is the safety and toxicology of the insecticide (Kra¨ mer and Mencke, 2001). Imidacloprid, 10% spot-on, was designed to offer a dermal treatment, which means it is applied externally onto a small dorsal area (a spot) of the animal’s skin. Criteria for the selection of an appropriate topical flea formulation are good solubility of the compound, good adhesion to the skin, good spreading properties, good local and systemic tolerance, stability, and compatibility with legal standards. The imidacloprid spot-on formulation, which meets all these requirements, contains 10 g a.i. in 100 ml nonaqueous solution. The efficacy of this formulation for flea control on cats and dogs has been reported (Kra¨ mer and Mencke, 2001). Imidacloprid applied at the target therapeutic dosage of 10.0 mg kg1 killed 99% of the fleas within 1 day of treatment, and continued to provide 99–100% control of further flea infestation for at least 4 weeks (Hopkins et al., 1996; Arther et al., 1997) (Figure 25). Studies using flea-infested cats (Jacobs et al., 1997b) proved that imidacloprid possess considerable potency against adult fleas on cats, and retains a high level of activity for 4–5 weeks. Imidacloprid was effective for both immediate relief from an existing flea burden (the therapeutic effect), and for longer-term flea control (the prevention or prophylactic effect) (Table 9). Imidacloprid spreads and acts using animal skin as the main carrier. The compound was shown to be localized in the waterresistant lipid layer of the skin surface, produced by sebaceous glands, and spread over the body surface and onto the hair (Mehlhorn et al., 1999). Insecticide spread over the skin surface was also reported from a clinical study that observed fast onset of flea control, as early as 6 h posttreatment (Everett et al., 2000). Therefore if the superficial fatty layer of

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Figure 25 Flea control achieved with imidacloprid 10% spot-on as confirmed by a dose-confirmation study in flea infested dogs (Hopkins et al., 1996).

Table 9 Geometric mean flea counts of two groups of cats, an untreated control and a group treated with imidacloprid 10% spot-on at a dosage of 10.0 mg kg1 bw at day 0 of the study After 1 day

After 2 days

Weeks after treatment

Control

Treated

Reduction (%)

Control

0 1 2 3 4 5 6

36.7 34.1 31.0 32.1 36.2 33.8 28.1

0.01 0.0 0.2 1.0 5.0 9.5 20.1

99.8 100.0 99.3 97.0 86.1 71.9 28.4

31.7

0.0

100.0

27.3 28.9 27.8 26.8 24.8

0.0 0.1 0.9 3.1 10.6

100.0 99.7 96.7 88.3 57.3

Treated

Reduction (%)

Reproduced from Jacobs, D.E., Hutchinson, M.J., Krieger, K.J., 1997b. Duration of activity of imidacloprid, a novel adulticide for flea control, against Ctenocephalides felis on cats. Vet. Rec. 140, 259–260.

the skin is removed by repeated swabbing with alcohol fleas consumed the same amount of blood in treated and untreated dogs. Thus imidacloprid localized in the lipid layer of the skin acts on adult fleas by contact. It was reported that imidacloprid is not taken up by the flea during blood feeding, but is absorbed via the flea’s smooth, nonsclerotized intersegmental membranes that are responsible for the insect’s mobility. Mehlhorn et al. (1999) concluded that this seems reasonable because of imidacloprid’s lipophilicity renders it incapable of passing through the sclerotized cuticle. Moreover, initial damage was seen in the ganglia close to the ventral body side (i.e., in subesophageal and thoracic ganglia). Fleas affected by imidacloprid treatment showed characteristic pathohistological changes (Mehlhorn et al., 1999). Muscle fibers and tissue around the subesophageal ganglion were damaged, with the mitochondria and axons showing intensive vacuolization (Figure 26). Imidacloprid’s mode of action corresponded with the structural findings (Mehlhorn et al., 1999), which showed overall destruction of

the mitochondria, damage of the nerve cells, and disintegration of the insect muscles (Figure 26). Imidacloprid’s activity on ectoparasitic insects results from its presence within the lipid layer of the host body surface. Since this lipid layer is always present, imidacloprid remains available for a prolonged time (Hopkins et al., 1996; Mehlhorn et al., 2001a), and reduces the likelihood of its removal during water exposure (Mehlhorn et al., 1999). The spectrum of imidacloprid activity to ectoparasites is not confined to fleas. It has also proven to be highly effective against both sucking lice (Linognathus setosus), and biting or chewing lice (Trichodectes canis) (Hanssen et al., 1999). Furthermore imidacloprid acted rapidly on all motile stages of sheep keds (Mehlhorn et al., 2001b). Sheep keds of the species Melophagus ovinus are wingless parasitic insects belonging to the dipteran family Hippobiscidae. Besides skin infection, which results in the loss wool quality and meat production, sheep keds are also known to transmit diseases, such as trypanosomiasis. However, ticks did not prove to

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Figure 27 Emergence of adult fleas from flea eggs incubated on blankets used by untreated (control) or imidacloprid treated cats. (Reproduced from Jacobs, D.E., Hutchinson, M.J., Ewald-Hamm, D., 2000. Inhibition of immature Ctenocephalides felis felis (Siphonaptera: Pulicidae) development in the immediate environment of cats treated with imidacloprid. J. Med. Entomol. 37, 228–230.)

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Figure 26 Transmission electron micrograph of a section through an adult cat flea exposed to imidacloprid for 1 h in vitro. Note the extensive damage at the level of muscle fibers and the subesophagal ganglion (vacuoles). CV, cellular cover of the ganglion; DA, degenerating axon; DC, degenerating nerve cell; DM, degenerating mitochondrion; FI, Fibrillar layer of connective tissue; Mu, Muscle fiber; TR, tracheole. Magnification 25 000. (Reproduced with permission from Mehlhorn, H., Mencke, N., Hansen, O., 1999. Effects of imidacloprid on adult and larval stages of the cat flea Ctenocephalides felis after in vivo and in vitro experiments: a light- and electronmicroscopy study. Parasitol. Res. 85, 625–637.)

be sensitive to complete control by imidacloprid (Young and Ryan, 1999). 5.3.10.1.2. Larvicidal activity Apart from the fastacting adulticidal activity of imidacloprid on flea populations, its larvicidal effects have also been investigated. Reinfestation of animals, from earlier deposited eggs, larvae, and preemerged adult fleas, can be overcome by the use of effective larvicidal compounds or insecticides acting on both the adult and the immature stages of fleas. In early studies using imidacloprid, larvicidal activity was observed in the immediate surrounding of treated dogs (Hopkins et al., 1997). Skin debris collected from treated dogs, when mixed into flea rearing media, showed high flea larva mortality. In repeated tests using the skin debris samples, collected at day 7 posttreatment, the larval mortality remained high at

100% even after 51 days (Arther et al., 1997). In in vitro studies, flea larvae survived for only 6 h when placed on clipped hair from imidacloprid treated dogs (Mehlhorn et al., 1999). Similar results have also been reported for cats (Jacobs et al., 2000). Adult flea emergence was reduced by 100%, 84%, 60%, and 74% in the first, second, third, and fourth week postimidacloprid treatment, in comparison to untreated controls (Figure 27). This persistent larvicidal activity is important, because in the absence of any larvicidal effect of an applied adulticide, reinfestation would occur from eggs deposited prior to treatment (Jacobs et al., 2001). Furthermore, cats wander freely outdoors, and thus may visit sites shared with other flea-infested domestic or wild animals. 5.3.10.1.3. Flea allergy dermatitis (FAD) FAD is a disease in which a hypersensitive state is produced in a host in response to the injection of antigenic material from flea salivary glands (Carlotti and Jacobs, 2000). Synonyms for FAD include flea bite allergy and flea bite hypersensitivity. In cats, the disease is also known as feline miliary dermatitis and feline eczema. FAD is one of the most frequent causes of skin conditions in small animals, and a major clinical entity in dogs. FAD is the commonest nonroutine reason for pet owners to seek veterinary advice. Hypersensitivity to flea bites is not only of importance to domestic pets, but is also an important cause of the common skin disease in humans, termed papular urticaria. Detailed investigations carried out on patients exposed to flea-infested pets have shown that the incidence of such reactions is quite high. Several field studies have been conducted, focusing on the efficacy of imidacloprid on cats and dogs with clinical signs of FAD (Kra¨ mer and Mencke, 2001). The efficacy of imidacloprid in flea removal,

Neonicotinoid Insecticides

and the resolution of FAD was tested in dogs and cats from single- and multiple-animal households (Genchi et al., 2000). Flea infestation was examined and FAD dermatitis lesions were ranked according to severity of typical clinical signs. Flea numbers dropped significantly after treatment of animals from both single- and multiple-animal households. In dogs clinical signs of FAD prior to treatment, decreased from 38% to 16% by day 14, and 6% by day 28, thus verifying a rapid adulticidal and high residual activity that lasted at least 4 weeks. There was an effective control of parasites, with rapid improvement of allergy until almost complete remission up to 28 days following the first application. Recently studies on the effect of imidacloprid on cats with clinical signs of FAD confirmed field data published by Genchi et al. (2000). Clinical signs of FAD, especially alopecia and pruritus were resolved after monthly treatment using imidacloprid (Keil et al., 2002) (Figure 28). Furthermore, controlling FAD is enhanced when blood feeding of fleas is reduced. This so-called ‘‘sublethal effect’’ or ‘‘antifeeding effect’’ was reported using very low concentrations of imidacloprid (Rust et al., 2001, 2002). 5.3.10.1.4. Imidacloprid as combination partner in veterinary medicinal products The ability of acaricides to repel or kill ticks, before they attach to a host and feed, is important for the prevention of transmission of tick born pathogens (Young et al., 2003). K9 AdvantixTM, an effective tick control agent (Spencer et al., 2003, Mehlhorn et al., 2003), is a spot-on product containing 8.8% (w/w) imidacloprid and 44% (w/w) permethrin. The mixture repels and kills four species of ticks, including Ixodes scapularis, for up to 4 weeks. It also repels and kills mosquitoes, and kills flea adults and larvae.

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Furthermore, a combination containing imidacloprid 10% (w/v) and permethrin 50% (w/v) in a spot-on formulation, has repellent and insecticidal efficacy, against the sand fly species (Phlebotomus papatasi) (Mencke et al., 2003), ticks (I. ricinus, Rhipicephalus sanguineus), and flea (C. felis felis) (Epe et al., 2003) on dogs. Another combination product, (Advantage HeartTM (10% w/v imidacloprid plus 1% w/v moxidectin), a macrolide antihelmintic, has been developed as a spot-on for dermal application to kittens and cats (Arther et al., 2003). It is intended for monthly application for control of flea infestations and intestinal nematodes, and for prevention of feline heartworm disease. It controls and treats not only established adult gastrointestinal parasites, but also developmental stages, including fourth instar larvae and immature adults of Toxocara cati in cats (Hellmann et al., 2003; Reinemeyer and Charles, 2003). Furthermore, the spot-on combination is safe and highly efficacious against T. canis and Ancylostomatidae in naturally infested dogs (Hellmann et al., 2003), as well as against Sarcoptes scabiei var. canis on dogs (Fourie et al., 2003).

5.3.11. Concluding Remarks and Prospects The discovery of neonicotinoids as a new class of nAChR ligands can be considered a milestone in insecticide research, and permits an understanding of the functional properties of insect nAChRs. Up to now the most current information regarding nAChRs originated from research with vertebrate receptors. The world market for insecticides is still dominated by compounds irreversibly inhibiting acetylcholinesterase, an important enzyme in the CNS of insects. The market share of these inhibitors,

Figure 28 Resolution of signs of flea allergy dermatitis (FAD) in cats treated with imidacloprid over a 84-day study period (Keil et al., 2002).

96 Neonicotinoid Insecticides

t0050

Table 10 Modes of action of the top selling 100 insecticides/ acaricides and their world market share (excluding fumigants, endotoxins and those insecticides with unknown mode of action) Market share (%) Mode of action

1987

1999

Change (%)

Acetylcholinesterase Voltage-gated sodium channel Acetylcholine receptor GABA-gated chlorine channel Chitin biosynthesis NADH dehydrogenase Uncouplers Octopamine receptor Ecdysone receptor

71 17 1.5 5.0 2.1 0 0 0.5 0

52 18 12 8.3 3.0 1.2 0.7 0.6 0.4

20.0 þ1.4 þ10.0 þ3.3 þ0.9 þ1.2 þ0.7 þ0.1 þ0.4

Reproduced with permission from Nauen, R., Bretschneider, T., 2002. New modes of action of insecticides. Pestic. Outlook 12, 241–245. ß The Royal Society of Chemistry.

and those insecticides acting on the voltage-gated sodium channel, account for approximately 70% of the world market (Nauen and Bretschneider, 2002) (Table 10). Today, the neonicotinoids are the fastest-growing group of insecticides (estimated marked share in 2005: about 15%), with widespread use in most countries in many agronomic cropping systems, especially against sucking pests but also against ectoparasitic insects. The relatively low risk and target specificity of the products, combined with their suitability for a range of application methods, also will ensure their success as important insecticides in integrated pest management (IPM) strategies.

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5.4 GABA Receptors of Insects S D Buckingham and D B Sattelle, University of Oxford, Oxford, UK ß 2005, Elsevier BV. All Rights Reserved.

5.4.1. Introduction 5.4.2. Ionotropic GABARs of Insects 5.4.2.1. Lessons from Vertebrate GABARs 5.4.2.2. Pharmacology of In Situ Insect GABARs 5.4.2.3. Diversity: Existence of Subtypes 5.4.2.4. Molecular Biology 5.4.2.5. Intracellular Modulation of GABARs 5.4.2.6. GABARs as Targets for Insecticides 5.4.2.7. Mechanisms of Insecticide Resistance 5.4.2.8. Molecular Basis of Selectivity of Insecticides for Insect GABARs 5.4.2.9. Single Channel Properties of Insect GABARs 5.4.2.10. Distribution 5.4.2.11. GABARs and Behavior 5.4.3. Metabotropic GABARs of Insects 5.4.3.1. A Cloned Insect Metabotropic (GABAB) Receptor 5.4.4. Conclusions

107 108 108 110 116 116 124 124 125 127 127 128 129 133 133 133

5.4.1. Introduction Receptors for the neurotransmitter g-aminobutyric acid (GABA) are abundant in the nervous systems of insects (Sattelle, 1990) where, as in vertebrates, they play a major role in inhibition. In 1985 when the first set of volumes of this series entitled Comprehensive Insect Physiology, Biochemistry, and Pharmacology was published, little was known about these proteins, despite the already established use of insect preparations to address fundamental questions in neurobiology, and despite the fact that GABA receptors (GABARs) were already considered to be possible insecticide targets. Since then, our understanding of insect GABARs has been transformed. Since 1985, studies on GABARs have benefited from the application of molecular biology (molecular cloning, functional expression, and determination of their gene structure), electrophysiology (singlechannel and whole-cell patch clamp recordings), integrative physiology (demonstrating a key role for such receptors in particular insect behaviors), genome analysis (yielding a complete set of ligand gated anion channels for the fruit fly, Drosophila melanogaster and the mosquito, Anopheles gambiae, and genetics (providing a detailed understanding of mutations that underly insecticide resistance). Radioligand binding studies have generated a detailed description of the pharmacology of insect GABAR binding sites. Electrophysiological studies, some using identified neurons, have contributed important information on

GABARs, both functional native and expressed recombinant receptors. Patch clamp methods, in particular, have provided key insights, enabling: (1) the analysis of single channel properties; and (2) description of the effects of chemicals on receptor kinetics. A number of studies have examined the distribution of GABARs and other molecular components of insect GABAergic synapses using antibodies to GABA, enzymes involved in GABA synthesis, and, more recently, antibodies to cloned GABAR subunits. Thus, arange of complementary experimental approaches have led to new discoveries on the roles of GABARs in sensory receptive field tuning, learning, and even a possible role in the insect circadian clock. A major advance in the study of insect GABARs resulted from the cloning of three candidate Drosophila ionotropic GABAR subunits, one of which resistant to DieLdrin(RDL) readily forms a functional, recombinant, homomeric GABA gated chloride channel. This was followed by the discovery of RDL orthologs in several insect orders. Thus, for the first time, it was possible to study in detail the physiology and pharmacology of an insect GABAR of known molecular composition. Consequently, in the years since the publication of the first edition of Comprehensive Insect Physiology, Biochemistry and Molecular Biology, there has been considerable growth in our understanding of insect GABARs (Hosie et al.,

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1997) and their vertebrate counterparts (Whiting, 2003). Insect GABAR subtypes were not well established in 1985, although even then data were beginning to emerge, to question earlier notions that benzodiazepine binding sites were not present in insects (Nielsen et al., 1978). Today, we know that insect GABARs rival the complexity of their vertebrate counterparts, and possess a number of sites for modulation by not only benzodiazepines, but also several other allosteric modulators. Again, in 1985, there was no evidence for the existence of an insect metabotropic receptor – today, three candidate insect metabotropic GABARs have been cloned and their characterization is now in progress. This chapter concentrates on new information that has accumulated since the previous edition of this series. After a brief review of findings up to 1985, we discuss the more recent findings on the physiology and pharmacology of insect ionotropic GABARs, examining each major chemical binding site in turn. Evidence for distinct subtypes of native insect GABARs is also discussed. Attention is then drawn to findings derived from the cloning and functional expression of recombinant insect GABARs, and their contributions to our understanding of (1) native receptors and (2) insecticide modes of action. Next, we consider the roles of ionotropic GABARs in insect behavior. Finally, new findings on metabotropic GABARs are described. By 1985, GABA was already known to be an inhibitory neurotransmitter at nerve–muscle junctions and in the central nervous system (CNS) of arthropods (Kuffler and Edwards, 1958; Usherwood and Grundfest, 1965; Otsuka et al., 1966), but very little is still known of insect GABARs. Most information was based on electrophysiology performed on native nerve and muscle receptors. However, GABA had been shown to induce hyperpolarizing responses in (mostly unidentified) central neurons of several insect species, including postsynaptic membranes of interneurons (Callec and Boistel, 1971; Callec, 1974; Hue et al., 1979) and unidentified cell bodies (Kerkut et al., 1969; Pitman and Kerkut, 1970; Walker et al., 1971; Roberts et al., 1981) in the terminal abdominal ganglion of the cockroach, Periplaneta americana. Dissociated cells of locust thoracic ganglia (Giles and Usherwood, 1985), as well as identified dorsal unpaired median neurons (DUMETi) that innervate the Extensor tibiae muscle of the embryonic grasshopper Schistocerca nitens (Goodman and Spitzer, 1980), were also found to respond to GABA with hyperpolarizations. Peripheral GABARs located on somatic muscle fibers had been described in some

considerable detail (Cull-Candy and Miledi, 1981; Cull-Candy, 1982). These early studies established that GABA mediates hyperpolarizing, largely inhibitory responses in insect neurons, but other than showing a sensitivity to picrotoxinin, little was known of the details of the pharmacology of these responses.

5.4.2. Ionotropic GABARs of Insects 5.4.2.1. Lessons from Vertebrate GABARs

Our understanding of both insect and vertebrate GABARs is based on binding studies, electrophysiology of in situ receptors and electrophysiology of expressed recombinant receptors, each approach offering its own particular advantages. Electrophysiological responses of neurons in insect or vertebrate nervous systems as well as cultured neurons provide a detailed account of the pharmacology, ion permeability, and kinetic properties of native GABARs. It is particularly convenient where identified neurons, or classes of neurons, are studied, as the analysis is restricted to a circumscribed (in some cases, perhaps homogeneous) class of receptors. Electrophysiological studies using heterologously expressed recombinant receptors offer the further advantage that the subunit composition of the receptors is known. Binding studies complement these functional approaches. However, when applied to native tissues the technique often pools data from multiple GABARs (as well as any other binding sites for the radioligand) from all cell types in the tissue under investigation. As functional and binding experiments yield different types of information, care must be exercised when comparing data obtained using these different techniques. For example, electrophysiological and binding studies typically address different states of the same receptor, which can also contribute to some of the apparent discrepancies. Nineteen ionotropic GABAR subunit molecules have been cloned from vertebrates (Whiting, 2003). They are members of the dicysteine-loop (cys-loop) family of ionotropic neurotransmitter receptors that also includes the nicotinic acetylcholine receptors (nAChRs) (Karlin, 2002), serotonin type 3 (5-HT3) receptors (5HT3Rs) (Reeves and Lummis, 2002), and strychnine sensitive glycine receptors (GlyRs) (Betz et al., 1999) of vertebrates as well as glutamate gated chloride channels (GluCls), a 5-HT gated chloride channel and a histamine gated chloride channel of invertebrates (review: Raymond and Sattelle, 2002). Each receptor molecule consists of five subunits arranged symmetrically around an integral, hydrophobic ion pore (Figure 1). Each polypeptide

GABA Receptors of Insects

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Figure 1 A schematic representation of the three-dimensional structure of ionotropic GABA receptors. Although more data are available for the structure of vertebrate ionotropic GABA receptors, it is probable that insect GABA receptors have a similar structure. (From Raymond, V., Sattelle, D.B., 2002. Novel animal-health drug targets from ligand-gated chloride channels. Nat. Rev. Drug Discov. 6, 427–436.)

subunit possesses a long N-terminal extracellular domain, which contains residues that contribute to the neurotransmitter (GABA) binding site and four transmembrane regions (M1–M4) (Figure 1). The second transmembrane domain (M2) contains most of the residues that line the chloride channel (Karlin and Akabas, 1995; Smith and Olsen, 1995). Vertebrate ionotropic GABARs may be divided into two pharmacological categories: GABAA receptors, which are all antagonized by the alkaloid, bicuculline, and regulated by a wide range of allosteric modulators such as benzodiazepines, barbiturates, and pregnane steroids; and GABAC receptors, which are insensitive to bicuculline as well as the majority of modulators of GABAA receptors (Bormann, 2000; Zhang et al., 2001). GABAC receptors are less liable to desensitize than GABAA receptors (Johnston, 1996). This physiological and pharmacological characterization is mirrored by corresponding structural differences between the two receptor subtypes, in that they are composed of distinct subunits. GABAA receptors are composed of subunits from several different classes (a1–6 b1–3 g1–3, d, e, p, and y) (Hedblom and Kirkness, 1997; Thompson et al., 1998; Neelands et al., 1999; Bonnert et al., 1999; Sinkkonen et al., 2000; Whiting, 2003), whereas GABAC receptors are believed to be composed of the three known isoforms of the r subunit, r1–3 (Sieghart, 1995; Djamgoz, 1995; McKernan and Whiting, 1996). GABAA receptors are complex allosteric proteins and contain, in addition to the agonist binding site, distinct sites for the actions of barbiturates, steroids, loreclezole, furosemide, picrotoxin (PTX), zinc, lanthanum, volatile

anaesthetics, benzodiazepines, and the anaesthetic, propofol. Although it should be noted that there is not perfect consensus on this division of vertebrate GABARs into A and C subtypes, this has been the view of the majority of workers to date and has received recent support (Bormann, 2000). This view will therefore be adopted for the purposes of this review. Functional expression studies, in which various combinations of subunits are heterologously expressed in Xenopus laevis oocytes or cell lines, have shown that the pharmacology of such recombinant vertebrate receptors is strongly influenced by subunit composition. For example, the specific subtype of b subunit present is a critical determinant of agonist pharmacology, whereas the a and g subunits are important in conferring sensitivity to benzodiazepines. When the a1 subtype is present in the heterologously expressed receptor it exhibits a pharmacology that resembles benzodiazepine type 1 pharmacology defined on the basis of in situ native receptor studies. Also, the inclusion of either the a2 or a3 subunit confers to the receptor the classical benzodiazepine type 2 pharmacology (Pritchett et al., 1989). The g2 subunit is essential for benzodiazepine activity (Pritchett et al., 1989), and the specific subtype of g subunit present in the receptor plays a major role in determining the receptor’s responsiveness to allosteric modulators, such as benzodiazepines. Taken together, experiments using recombinant expression have suggested that native vertebrate GABAA receptors are composed of at least a, b, and g subunits, a conclusion supported by several immunoprecipitation studies

110 GABA Receptors of Insects

(Mohler et al., 1992; Khan et al., 1994; Araujo et al., 1996; Jechlinger et al., 1998; Nusser et al., 1999). About 35% of GABAA receptors in the brain contain a1b2g2 subunits, 20% contain a3b3g2, and 15% contain a2b3g2 subunits. Other combinations are present, but they are less common (Whiting, 2003). With fewer laboratories working in the field the growth in our knowledge of insect GABARs has been slower than that of their vertebrate counterparts. It is clear, however, that while all vertebrate and insect ionotropic GABARs are blocked by the plant derived toxin, picrotoxinin, a number of pharmacological differences distinguish insect GABARs from vertebrate GABAA and GABAC receptors. For example, insect ionotropic GABARs do not readily fit the vertebrate classification. There are two principal differences. First, the majority of insect GABARs, unlike GABAA receptors, are insensitive to bicuculline, although bicuculline sensitive insect GABARs do exist. Secondly, most insect GABARs are pharmacologically distinct from GABAC receptors in that they are subject to allosteric modulation by benzodiazepines and barbiturates (Sattelle, 1990). There are other minor differences in aspects of their detailed pharmacology, each of which will be addressed in the following sections. 5.4.2.2. Pharmacology of In Situ Insect GABARs

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5.4.2.2.1. Agonist site Early binding studies on cockroach (P. americana) nerve cord preparations using [3H]GABA and [3H]muscimol indicated that insect GABARs, unlike vertebrate GABAA receptors, were insensitive to 3-aminopropanesulfonic acid (3APS). Also, muscimol is of the same, or higher, potency as GABA on insect GABARs (Lummis and Sattelle, 1985, 1986). The most potent displacers of [3H]GABA binding to cockroach nerve cord preparations were found to be GABA itself and muscimol, with IC50s of 130 and 710 nM respectively. In contrast, the other vertebrate agonists tested (thiomuscimol, 4,5,6,7-tetrahydroisoxazolo[5,4-c] pyridin-3-ol (THIP), 3-APS, and isoguvacine) had IC50s greater than 30 mM (Lummis and Sattelle, 1986). The dissociation rate for GABA binding was estimated to be around 0.116 min
1 and the estimated association rate was approximately 316 mM
1 min
1. The molecular structures of some GABAR agonists are illustrated in Figure 2. Later, electrophysiological studies were performed on GABARs in an identified Periplaneta motor neuron (the fast coxal depressor, Df) (Sattelle et al., 1988), and on cloned Drosophila GABARs containing only RDL subunits heterologously expressed in oocytes or insect cell lines (Millar et al., 1994; Hosie and

Figure 2 The molecular structures of some common agonists of vertebrate and insect ionotropic GABA receptors.

Sattelle, 1996a, 1996b; Buckingham et al., 1996). The only major difference between the [3H]GABA displacement profile and the functional studies is that isoguvacine is a potent agonist on native, functional insect GABARs (Sattelle et al., 1988), providing a further distinction from vertebrate GABARs. The reason for this apparent discrepancy between physiology and binding data is unclear, but may be attributable to the fact that the two methods probe different receptor states, and that the two approaches may probe different pools of receptor subtypes. The sensitivity of insect GABARs to ZAPA ((Z)-3-[(aminoiminomethyl)thio]prop-2-enoic acid) (Taylor et al., 1993) also distinguishes them from vertebrate GABAA receptors, which are largely insensitive to this compound (Woodward et al., 1993; Qian and Dowling, 1993). Reports from several laboratories show a lack of action of established vertebrate GABAB agonists or antagonists on insect GABARs (Lees et al., 1987; Sattelle et al., 1988; Murphy and Wann, 1988; Bermudez et al., 1991; Wolff and Wingate, 1998). However, this does not preclude the existence of GABAB receptors with unusual pharmacology. For example, the GABABR agonist baclofen and the GABAB antagonist saclofen were inactive on GABARs of Df (Sattelle et al., 1988). However, more recent experiments have identified GABAB-like responses, which differ significantly in their pharmacology from vertebrate GABAB receptors in being insensitive to baclofen (Bai and Sattelle, 1995). The presence of these insect GABAB receptors was only detected when responses mediated through ionotropic GABARs were blocked by PTX.

GABA Receptors of Insects

Most insect ionotropic GABARs are relatively insensitive to the GABA analog CACA (cisaminocrotonic acid), which is an agonist of vertebrate GABAC, but not GABAA, receptors. For example, visual interneurons in the fly, Calliphora erythrocephala, are CACA insensitive (Brotz and Borst, 1996). However, it should be noted that CACA activates GABARs that mediate inhibition by identified filiform hair receptors of an identified projection interneuron, a circuit that mediates wind elicited responses (Gauglitz and Pfluger, 2001). Taurine (2-aminoethanesulfonic acid), an abundant amino acid in both vertebrates and insects, may act as a coagonist with GABA in certain insects. It is abundant in the nervous system of the locust, Schistocerca gregaria (Whitton et al., 1987), where its distribution changes following periods of intense muscular activity. When ionophoretically applied to isolated locust neurons in culture, taurine evokes membrane conductance increases very similar to those evoked by GABA (Whitton et al., 1994). Indeed, responses to both GABA and taurine are both blocked by PTX and enhanced by flunitrazepam, consistent with their acting through the same receptor. It was therefore suggested that taurine may act at the same site as GABA, serving to modulate the GABA response. In the migratory locust, Locusta migratoria, taurine is coexpressed with GABA in local and intersegmental inhibitory interneurons but not in efferent neurosecretory or paracrine cells, suggesting a role in preventing hyperexcitation during stressful conditions (Stevenson, 1999). 5.4.2.2.2. Competitive antagonists A consistent feature of most, but not all, insect GABARs is that they are insensitive to the plant derived alkaloid, bicuculline (Figure 3), a defining blocker of vertebrate GABAA receptors (Curtis et al., 1971; Watson and Girdlestone, 1995). Extrasynaptic GABARs on the motor neuron, Df (Sattelle et al., 1988; Buckingham et al., 1994), and synaptic GABARs on the giant interneuron 2 of P. americana (Buckingham et al., 1994) are insensitive to bicuculline, as are many locust neuron ionotropic GABARs (Benson, 1988; Lees et al., 1987). Bicuculline does not block either the cloned Drosophila GABAR, RDL (Millar et al., 1994), or the cloned Heliothis virescens RDLlike GABAR (Wolff and Wingate, 1998). GABA mediated inhibitory postsynaptic potentials (IPSPs) recorded in an identified locust (S. gregaria) interneuron are also bicuculline insensitive (Watson and Burrows, 1987). Similarly, bicuculline insensitivity is a feature of GABARs that mediate inhibition by identified filiform hair receptors of an identified projection interneuron, a circuit that mediates wind

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Figure 3 The molecular structures of some common convulsant antagonists of vertebrate and insect ionotropic GABA receptors.

elicited responses (Gauglitz and Pfluger, 2001). Responses to muscimol in motion sensitive visual interneurons of the fly, C. erythrocephala, are also insensitive to bicuculline (Brotz and Borst, 1996). Furthermore, insensitivity to bicuculline is also seen for certain spider GABARs (Panek et al., 2002). However, there is a body of evidence that bicuculline does block ionotropic GABARs of some adult Manduca sexta abdominal ganglion neurons (Roberts et al., 1981; Waldrop et al., 1987; Waldrop 1994) as well as electrically or odor evoked IPSPs and GABA responses in projection neurons of antennal lobes of this species (Christensen et al., 1998). Recently, Sattelle et al. (2003) have also shown that bicuculline insensitive GABARs are present in the larval nervous system of M. sexta. Further, the observation that bicuculline is effective in recovering behaviors disrupted by the systemic administration of GABA transporter blockers (Leal and Neckameyer, 2002) suggests that, despite the lack of bicuculline action on RDL, bicuculline

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may nonetheless block some D. melanogaster GABARs. In addition, bicuculline also blocks certain nicotinic acetylcholine receptors (nAChRs) of insects (Benson, 1988; Buckingham et al., 1994) and vertebrates (Rothlin et al., 1999). One attempt to resolve the issue of bicuculline sensitivity tested the possibility that such divergent reports may reflect pharmacological differences between extrasynaptic and synaptic GABARs (Buckingham et al., 1994). The cockroach, P. americana, offers a convenient model for such an approach because of the number of indentified neurons with ready access to both their cell bodies and their synapses. Motor neuron Df has a population of well-characterized GABARs on its cell body membrane, and giant interneuron 2 (GI 2) not only presents an accessible cell body, but also receives synaptic inputs from nerve X. IPSPs measured in GI 2 using the oil gap recording method in response to electrical stimulation of nerve X were blocked by both bicuculline (100 mM) and PTX (0.01 mM). However, bicuculline injected directly into the neuropil in the region of these synapses did not block IPSPs. The apparent block of synaptic GABARs by bicuculline was found to be due to inhibition of nAChRs upstream in the synaptic pathway. While the study failed to find differences in the pharmacology of synaptic and extrasynaptic GABARs, it did show that three separate populations of GABARs exist: synaptic receptors in the neuropilar branches of GI 2, extrasynaptic GABARs on the cell body of GI 2, and extrasynaptic GABARs on Df, all of which are bicuculline insensitive. The finding that bicuculline acts on nAChRs does not, however, explain the block of IPSPs by bicuculline in M. sexta interneurons (Waldrop et al., 1987), since ACh responses in this preparation are insensitive to direct bicuculline application (Waldrop and Hildebrand, 1988). Thus, there is some crossover between the pharmacophore for ligand binding sites of insect ionotropic GABARs and nAChRs. Whereas most insect GABARs are insensitive to bicuculline, many insect ACh receptors, in contrast, are blocked by bicuculline (Buckingham et al., 1994; Waldrop, 1994). At the same time, imidacloprid, an insecticide acting as a partial agonist of nAChRs, can also partially block GABARs of honeybee Kenyon cells, albeit at relatively high concentrations (Deglise et al., 2002). In addition to their abundant expression in the CNS, and in contrast to vertebrates, insect GABARs are also expressed on muscle fibers. In many respects, these muscle insect GABARs resemble those of the nervous system in that they are sensitive to muscimol and to the GABA channel blocker, picrotoxinin (Scott and Duce, 1987; Murphy and Wann, 1988).

However, until 1997, no detailed comparison had been reported for muscle and neuronal GABARs in the same animal for any insect species. One study, therefore, set out directly to compare GABARs on fibers of the coxal levator muscle (182c,d) of the cockroach, P. americana (Schnee et al., 1997), with the well-characterized GABARs on the motor neuron, Df, of the same species (Sattelle et al., 1988). Both were sensitive to PTX and insensitive to bicuculline, but there were significant pharmacological differences between them. Unlike the neuronal receptors, the muscle GABARs were insensitive to isoguvacine. The EC50 of muscimol relative to GABA was also much lower for muscle than for Df. The muscle GABARs were also sensitive to certain cyclodienes: 12-ketoendrin, endrin, and heptachlor epoxide all reduced GABA responses with IC50s in the 1–10 nM range. Similarly, micromolar concentrations of the cyclodiene, heptachlor, produced a complete block of muscle receptors, whereas concentrations as high as 100 mM resulted in about 50% block of GABARs on cockroach motor neuron Df (Lummis et al., 1990). Micromolar tert-butyl-bicyclo[2.2.2]phosbrothionate (TBPS) and tert-butyl-bicylo orthobenzoate (TBOB) were almost without effect on muscle receptors. Hence, in the cockroach there are some clear differences between neuronal and muscle GABARs. The observation that muscle GABARs are significantly more sensitive to cyclodienes than the GABARs of motor neuron Df is interesting in view of the fact that mutants of RDL are resistant to cyclodienes, yet RDL is expressed exclusively in the nervous system. This discrepancy remains to be addressed. 5.4.2.2.3. Noncompetitive antagonists and convulsants Several convulsants (Figure 3) are noncompetitive antagonists of insect GABARs and this site is the target of many insecticides, including the cyclodienes (such as endrin, dieldrin), hexachlorocyclohexanes (such as lindane), and bicycloorthobenzoates (such as 1-(4-ethynylphenyl)4-n-propyl-2,6, 7-trioxabicyclo[2.2.2]octane (EBOB)) (Matsumura and Ghiasuddin, 1983; Lummis and Sattelle, 1986; Wafford et al., 1989; Lummis et al., 1990; Deng et al., 1993; Anthony et al., 1994). The earliest evidence that cyclodienes act at the convulsant site came from binding studies using [3H]dihydropicrotoxinin, which indicated that the cyclodiene and PTX binding sites were similar (Matsumura and Ghiasuddin, 1983). Further, houseflies resistant to dieldrin show cross-resistance to many polychlorocycloalkanes and phenylpyrazoles, and to some bicycloorthobenzoates, but not to avermectins

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(Deng et al., 1991; Deng and Casida 1992; Cole et al., 1993; Casida, 1993). Initial studies using labeled probes for the convulsant site were hampered by the limitations of [3H]dihydropicrotoxinin as a probe. These were largely overcome as improved radioligands became available. Specific, saturable binding of [35S]TBPS to cockroach nerve cord membranes yielded a Kd of around 18 nM and Bmax of around 177 fmol mg
1 protein (Lummis and Sattelle, 1986). This binding was inhibited by GABA and dihydroavermectin B1a at micromolar concentrations, suggesting the presence of an allosteric linkage between these sites in insect GABARs. Radioligand binding and toxicity approaches used by John Casida’s laboratory have been particularly fruitful not only in identifying the sites of action of many insecticidally relevant compounds, but also in providing a pharmacological dissection of the convulsant site. The [3H]EBOB binding site of housefly heads was found to be identical to, or to overlap with, a wide array of structurally distinct insecticides (Deng et al., 1991) including cyclodienes, bicycloorthobenzoates, and PTX, but not avermectins. This was pursued further in a comprehensive study of the convulsant site examining the displacement of labeled EBOB and TBPS by 29 different compounds (Deng et al., 1993). Binding of [35S]TBPS to housefly head membranes had a Kd of 145 nM and Bmax of 2.4 pmol mg
1 protein, and both [35S]TBPS and [3H]EBOB were displaced by a number of polychlorocycloalkanes, bicycloorthobenzoates, and phenylpyrazoles. The binding parameters resembled those of vertebrate tissues with respect to pH dependence, anion specificity and kinetics of association, and dissociation. Comparing the modes of competition of each compound class (i.e., competitive versus noncompetitive) and their relevance to toxicity, this study concluded that four subsites can be defined pharmacologically: site A (the EBOB site) overlaps with site C (the phenylpyrazole site) and with site B (the TBPS site), but sites B and C do not overlap. The avermectin site (site D) is distinct from all of the other three sites, but is allosterically linked to sites A and C. Similar studies added weight to the conclusion that insect GABARs differ significantly from their vertebrate counterparts at the noncompetitive binding sites. For instance, they are significantly less sensitive to the polychlorocycloalkane, cage convulsants (such as TBPS and TBOB), and PTX, compounds that by contrast are potent inhibitors of vertebrate GABARs (Rauh et al., 1990). A single exception to this rule is furnished by the polychlorocyclohexane insecticide, heptachlor, and its epoxide

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metabolite, which more potently displaced labeled TBPS from Musca domestica membranes than from rat (Lummis et al., 1990). However, when muscle GABARs of the cockroach, P. americana, were compared to those on an identified motor neuron of the same species (Schnee et al., 1997), muscle GABARs were found to be more sensitive to lindane and to heptachlor and its epoxide metabolite. One fertile approach to understanding the convulsant site has been to compare the efficacies of a systematically varying set of PTX analogs in either blocking the receptor or in displacing radiolabeled probes. Such studies on vertebrate tissues led to the proposal (Ozoe and Matsumura, 1986) that the key elements in determining picrotoxane activity are the presence of a bulky lipophilic group and at least two electronegative centers. Several studies have emphasized the importance of two groups in the PTX molecule (the hydroxyl and the isopropenyl groups) in conferring high lethality (Jarboe et al., 1968), on displacing [3H]-dihydropicrotoxin (Ticku et al., 1978), and as well as affecting electrophysiological responses of in situ (Kudo et al., 1984) and recombinant GABARs (Anthony et al., 1993). These conclusions were drawn in particular from the observation that fluoropicrotoxinin, which substitutes a fluorine at the hydroxyl position, is almost equally active as the parent molecule, whereas PTX acetate is much reduced in potency, and that picrotin, which adds a hydroxyl group to the isopropenyl, is also comparatively ineffective. Such findings suggest that for vertebrates, the hydroxyl group may act as a hydrogen bond acceptor, that substitutions at the hydroxyl group are liable to steric hindrance, and that the isopropenyl group must be lipophilic or electron-rich (Anthony et al., 1993) for maximal activity. To determine whether the same is true for insect GABARs, one study (Anthony et al., 1994) examined the actions of a series of PTX analogs on the well-defined GABARs on the identified motor neuron, Df, of the cockroach, P. americana. The rank order of potency was the same as for expressed chick optic lobe GABARs, suggesting that the convulsant sites of insect and vertebrate GABAA receptors share certain common key features. Similar findings were obtained for RDL, a cloned insect GABAR (see Section 5.4.2.4.3.2). The reduced sensitivity of insect GABARs to TBPS slowed the development of our understanding of insect convulsant site probes. Recently, however, two compounds have been deployed with selectivity for insect over vertebrate GABARs, BIDN, and fipronil (Figures 3 and 4). BIDN ([3H]3,3-bis-trifluoromethyl-bicyclo[2,2,1] heptane-2,2-dicarbonitrile) was developed using a

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combined biochemical and pharmacophore modeling approach (Rauh et al., 1997a, 1997b). A large library of compounds was systematically screened for their ability to displace [35S]TBPS from rat brain membranes. Several bicyclic dinitriles were identified and further screened for their selectiviy for insect GABARs over rat GABARs. This approach identified the common structural features of known convulsant site ligands, which helped to determine the key pharmacophore features (Figure 5) This was done by manually aligning each of a series of polycyclic dinitriles over a representation of the PTX molecule using the Sybyl software package (SYBYL Molecular Modeling Software, Tripos Associates, Inc., St. Louis, MO). Key pharmacophore elements identified were the presence of a polarizable moiety separated by a fixed distance from two hydrogen bond-accepting (i.e., electronegative) elements (Figure 5), confirming and extending the model proposed earlier by Ozoe and Matsumura (1986). A number of new potential ligands were developed from this pharmacophore model. Of these, the most useful was BIDN (Rauh et al., 1997a, 1997b).

Figure 4 The molecular structures of two potent antagonists of insect ionotropic GABA receptors, dieldrin and fipronil, both of which have been used as insecticides.

Binding of [3H]BIDN to southern corn rootworm (Diabrotica undecimpunctata) membranes was saturable and specific with high affinity (26 nM) and a Hill coefficient of unity suggesting that BIDN binds to a single class of receptor sites (Rauh et al., 1997b). Such high-affinity, specific binding was confirmed in the same study for five other insect species, including P. americana and M. domestica, whereas binding of BIDN to rat membranes was of lower affinity (around 250 nM). The same study examined the actions of BIDN using electrophysiological techniques, which showed that BIDN blocks GABARs of the identified cockroach motor neuron Df as well as those present on the coxal levator muscles of the same species. BIDN also blocks GABA evoked depolarizations of thoracic neurons in the southern corn rootworm, (Rauh et al., 1997b). The estimated IC50 for BIDN action against central (on the cell body of motor neuron, Df) and peripheral (on coxal levator muscle fibers) GABARs of P. americana was 0.9 mM and 2.5 mM, respectively. Although showing a degree of selectivity for insect over vertebrate receptors, the vertebrate toxicity of BIDN was a factor precluding its further development as a commercial pesticide. Another insect GABAR ligand is the potent, widely used insecticide active on insect GABARs, the phenyl pyrazole, fipronil. Fipronil is a potent antagonist of cloned insect GABARs (Hosie et al., 1997; Grolleau and Sattelle, 2000) and has also been used as the basis for the development of a photo-affinity probe for the convulsant site (Sirisoma et al., 2001). There is abundant evidence that BIDN and fipronil act at the convulsant site. Fipronil competes for EBOB binding to housefly head membranes (Ratra and Casida, 2001). BIDN (Hosie et al., 1995b), picrotoxinin (Shirai et al., 1995), and fipronil (Hosie

Figure 5 The ideal pharmacophore for the picrotoxin molecule. This was derived from a number of structure–function studies that have revealed the structural features that confer potency to picrotoxin analogs. This was used to predict that a series of bicyclic dinitriles would have potency at the picrotoxinin site. (From Rauh, J.J., Holyoke, C.W., Kleier, D.A., Presnail, J.K., Benner, E.A., et al., 1997b. Polycyclic dinitriles: a novel class of potent GABAergic insecticides provides a new radioligand, [3H]BIDN. Invert. Neurosci. 3, 261–268.)

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et al., 1995b) all block RDL (see below) in a voltage independent, partly competitive and partly noncompetitive manner. However, although dieldrin displaces [3H]BIDN binding competitively, PTX does not (Sattelle et al., 1995), suggesting that the PTX and the dieldrin site are not identical. This confirms an earlier report (Deng et al., 1993) that identified no less than four convulsant antagonist binding sites in the housefly GABAR that can be differentiated by [3H]EBOB and [35S]TBPS. It is therefore likely that BIDN, fipronil, EBOB, and PTX act at distinct but overlapping sites. Together, these compounds therefore provide a powerful tool-kit with which to further dissect the actions of insecticides on GABARs. Single channel properties of RDL expressed in a Drosophila cell line suggested that BIDN and fipronil both reduce the mean open time of GABAR chloride channels, but do so by acting at distinct sites (Grolleau and Sattelle, 2000) – see Section 5.4.2.4.3.2. There are no doubt a number of naturally occurring ligands with GABAergic actions that await discovery. An interesting class of compounds acting at this site is the a and b thujones, the active ingredients of absinthe and present in certain herbal medicines. These compounds act at the convulsant site to block GABARs noncompetitively, and have insecticidal activity (Hold et al., 2000, 2001). It will be of interest to study possible actions of other plant derived compounds on insect GABARs. 5.4.2.2.4. Allosteric modulators 5.4.2.2.4.1. Benzodiazepines In 1985, there was still some controversy as to whether insect nervous tissues possess receptors for benzodiazepines. One report suggested a lack of benzodiazepine receptors in invertebrates (Neilsen et al., 1978) but this was followed by a number of reports (Abalis et al., 1983; Lummis and Sattelle, 1986; Robinson et al., 1985) of sensitivity of insect GABARs to benzodiazepines. All these reports indicated that the pharmacology of the insect benzodiazepine site differs from most of its vertebrate counterparts. Specific [3H]flunitrazepam binding to cockroach (P. americana) nerve cord membranes yielded an estimated Kd of 383 nM and Bmax of 5.5 pmol mg
1 protein (Lummis and Sattelle, 1986). The most potent displacer of this binding was Ro5-4864 (IC50 ¼ 1 mM), which was only slightly more potent than flunitrazepam (IC50 ¼ 1.6 mM). Clonezapam was much less effective (IC50 ¼ 25 mM). The pharmacological profile of the [3H]flunitrazepam binding site in P. americana was Ro5-4864 (IC50 ¼ 0.6 mM) > diazepam (IC50 ¼ 1.0 mM) > flunitrazepam (IC50 ¼ 1.6 mM)  clonazepam (IC50 ¼ 25 mM)  Ro15-1788 (IC50 ¼ 100 mM). Allosteric linkage between the agonist and benzodiazepine

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sites was confirmed by the fact that 0.1 mm GABA enhanced flunitrazepam binding by some 150%. Furthermore, flunitrazepam enhanced responses to GABA in locust neuronal somata (Lees et al., 1987; Whitton et al., 1994) and in the motor neuron Df of the cockroach at micromolar concentrations (Sattelle et al., 1988). The mode of action of benzodiazepines seems to be, as in vertebrates, an increase in the frequency of opening of the GABA gated channel (Shimahara et al., 1987). The potent displacement of [3H]flunitrazepam binding in cockroach by Ro5-4864 and its potentiation of GABA responses suggests that cockroach CNS benzodiazepine receptors resemble vertebrate peripheral benzodiazepine binding sites rather than central ones (Lummis and Sattelle, 1986). These peripheral binding sites in vertebrates are located principally in the kidney and liver, where they are associated with calcium channels. Nevertheless, the observation that flunitrazepam and GABA binding are allosterically linked, along with physiological evidence discussed below, confirm that the [3H]flunitrazepam binding site in insect CNS is indeed part of a GABAR. Flunitrazepam and diazepam have no effect upon GABARs of coxal levator muscle of cockroach, even at concentrations of up to 100 mM (Schnee et al., 1997) or upon the cloned H. virescens GABAR (Wolff and Wingate, 1998). Flunitrazepam does, however, evoke responses to GABA in the cockroach Df motor neuron (Sattelle et al., 1988) and other neuronal preparations (review: Sattelle, 1990). That there are differing accounts of the actions of benzodiazepines upon insect GABARs is not surprising, given the great diversity in benzodiazepine pharmacology of vertebrate GABARs and the acute sensitivity of its pharmacology on subunit composition. A sensitivity to Ro5-4864 is probably the most significant distinguishing feature since the site of action of this compound in vertebrates is not on GABARs. Further, the lack of evidence for actions of benzodiazepines on insect muscle GABARs may represent an additional distinction between muscle and neuronal GABAR subtypes in insects, although more benzodiazepines would have to be tested to confirm this hypothesis. 5.4.2.2.4.2. Barbiturates Although there have been relatively few studies, there is evidence that insect GABARs possess a site for the action of barbiturates. Enhancement of responses to GABA recorded in locust neurons by barbiturates has been reported (Lees et al., 1987), and phenobarbital enhances responses mediated by the heterologously expressed H. virescens GABAR (Wolff and Wingate, 1998). In contrast, sodium pentobarbital at 100 mM failed to modify

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responses to GABA recorded in cockroach coxal levator muscle (Schnee et al., 1997). 5.4.2.2.4.3. Other allosteric modulators In addition to the benzodiazepine and barbiturate allosteric modulator sites, there are reports of modulatory actions of other compounds. Pregnane steroids displace [35S]TBPS binding from M. domestica membranes, but with low affinity, while the insect steroids ecdysone and 20-hydroxyecdysone are almost without effect (Rauh et al., 1990). Other allosteric modulators of insect GABARs include thymol (Gonzalez-Coloma et al., 2002). Priestley et al. (2003) have shown that thymol acts at a novel allosteric site. Propofol, widely used as an anaesthetic, enhances GABA responses mediated through RDL by up to about nine-fold with an EC50 of about 8 mM (Pistis et al., 1996a). The action of propofol on vertebrate GABAA receptors is critically dependent upon the presence of a specific amino acid in the second transmembrane domain of the receptor (Pistis et al., 1999). 5.4.2.3. Diversity: Existence of Subtypes

One of the most convincing demonstrations of the existence of multiple GABAR subtypes is provided where different tissues within the same animal possess receptors of distinct pharmacology. Bicuculline sensitive and insensitive GABARs are present in the nervous system of the moth, M. sexta (Waldrop, 1994; Waldrop et al., 1987a,b; Sattelle et al., 2003). In P. americana, muscle receptors differ from nervous system receptors in their insensitivity to isoguvacine and higher sensitivity to channel blockers, along with differences in their sensitivity to barbiturates and benzodiazepines, although the latter is less likely to be useful in defining the muscle subtype given the wide variation in benzodiazepine pharmacology among different GABARs. Taken together, however, the differences in pharmacology between insect muscle and neuronal receptors is sufficient to justify considering them as distinct receptor subtypes. An even more convincing demonstration of the existence of GABAR subtypes is furnished where two types of GABARs can be demonstrated on the same cells. Here, work on identified cells is particularly compelling. DUM (dorsal unpaired median) neurons of the cockroach have been shown to possess at least two subtypes of ionotropic GABAR, one of which is regulated by intracellular calcium ions through a calmodulin pathway, while the other is sensitive to CACA but seemingly not regulated by intracellular calcium (Alix et al., 2002).

Clearly, not only are insect GABARs to be distinguished pharmacologically from the major vertebrate subtypes, but there is also considerable evidence to suggest that insect GABARs are not of one pharmacologically uniform type. However, there is not enough data available yet to group all the known insect GABARs into a clearly delimited classification scheme. We would argue, however, that a classification into bicuculline sensitive versus insensitive and a subset of bicuculline insensitive muscle receptors is justified on the basis of currently available evidence. 5.4.2.4. Molecular Biology

5.4.2.4.1. Cloned receptors Ionotropic GABAR subunits have been cloned from eight species of insect. These fall into three distinct types: RDL, which has been found in a number of insect species, and lignad-gated chloride channel 3 (LCCH3) and GABAA and glycine receptor-like subunit of Drosophila (GRD), which are only known to date in Drosophila. All three are structurally similar to vertebrate GABAA receptor subtypes, and more generally to the cys-loop family of neurotransmitter receptors: they have four transmembrane domains of which the second possesses an amphipathic helix and a dicysteine motif in the extracellular domain (see Hosie et al., 1997). GRD resembles both ionotropic GABA and glycine receptors of vertebrates (Harvey et al., 1994). It has 33–44% identity to vertebrate GABAR subunits, but is distinguished by a large insertion between the dicysteine loop and the first transmembrane spanning region. Such a feature is not seen in any other ligand gated chloride channel. Its highest sequence identity is to vertebrate GABAA a subunits (40–44% identity), but it has almost the same degree of sequence identity to vertebrate glycine receptor a subunits (40–41% identity). GRD does not form receptors in Xenopus oocytes, but its presumed M2 region contains seven of the eight amino acid motif (TTVLTMTT) that is a signature of ligand gated chloride channels (Harvey et al., 1994). Furthermore, Xenopus oocytes, injected with mRNA encoding the GRD subunit, do not respond to a wide range of amino acids and glutamate receptor agonists, neither does GRD form functioning receptors when coinjected with bovine GABAA a1 or b1 subunit or rat glycine b subunits (Harvey et al., 1994). In light of the failure of GRD to support GABA responses, it is probably significant that GRD lacks any equivalent to the ligand binding domain II (TGSY) of GABAA receptors. There is, therefore, reason to suspect that GRD may not even be a GABAR subunit. LCCH3 has

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high (approximately 47%) identity to vertebrate GABAA b subunits (Henderson et al., 1993, 1994). 5.4.2.4.2. RDL: a model insect ionotropic GABAR The rdl (resistance to dieldrin) gene was originally isolated from a naturally occurring dieldrin resistant strain of D. melanogaster (ffrenchConstant et al., 1991, 1993a). The rdl locus maps to position 66F of chromosome III. It has nine exons, alternative splicing of two of which gives rise to four gene products (Figure 6), of which the most extensively studied are RDLac and RDLdb. The rdl gene is not limited to Drosophila, nor indeed even to the Diptera. RDL-like GABAR subunits with 85–99% amino acid identity have also been cloned from Myzus persicae (Anthony et al., 1998), H. virescens (Wolff and Wingate, 1998), Aedes aegypti (Thompson et al., 1993a), Drosophila simulans (ffrench-Constant et al., 1993a), M. domestica (Thompson et al., 1993a), Blatella germanica

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(Thompson et al., 1993b; Kaku and Matsumura, 1994), Tribolium castaneum (Miyazaki et al., 1995), and Hypothenemus hampei (Andreev et al., 1994), indicating that it is widespread throughout insect orders. The RDL polypeptide has between 30% and 38% identity with vertebrate GABAR subunits (about the same identity among different vertebrate subunit types). RDL GABAR subunits encoded by the rdlgene are distributed throughout the adult (Harrison et al., 1996) and embryonic (Aronstein and ffrenchConstant, 1995) nervous system of Drosophila, particularly in the optic lobes, ellipsoid body, fanshaped body, ventrolateral cerebrum, and glomeruli of antennal lobes. RDL is not expressed in muscle. Furthermore, in Drosophila, RDL expression is largely confined to the neuropil, with no discernible expression on neuronal cell body membranes or muscle (Harrison et al., 1996), although cell bodies were clearly stained in P. americana (Sattelle et al.,

Figure 6 The locations of the determinants of agonist potency of cys-loop receptors are highly conserved. In this schematic aligmnent of cys-loop receptor subunits, the location of the cysteine-loop is indicated by the line above the subunit. The location of known determinants of agonist potency in nACh receptors, termed loops A–E, are marked yellow, while those of benzodiazepine potency are marked green. The exon boundaries of Rdl-encoded polypeptides are marked by vertical red lines. As a result of the alternative splicing of two exons, the Drosophila Rdl gene encodes four polypeptides, each of which exhibits characteristic features of GABAA receptor subunits. The alternatively spliced exons (3 and 6) encode regions of the extracellular N-terminal domain. There are two variant forms of each exon; those of exon 3 are termed ‘‘a’’ and ‘‘b’’ and differ by two residues, while the alternate forms of exon 6, termed ‘‘c’’ and ‘‘d’’, differ at 10 residues. Thus, depending on the splice variants present in a given polypeptide, the different Rdl encoded subunits may be referred to as RDLac, RDLbd, etc. mRNAs encoding all four splice variants have been identified in embryonic D. melanogaster. The positions of these variant residues are marked in purple. The two alternate residues encoded by exon 3 lie close to a known determinant of the agonist potency of GABA receptors, while the 10 variant residues encoded by splice variants of exon 6 span a region that is poorly conserved in vertebrate GABA receptor subunits, but which corresponds to determinants of agonist potency in nAChRs. (Reproduced with permission from Hosie, A.M., Aronstein, K., Sattelle, D.B., ffrench-Constant, R.H., 1997. Molecular biology of insect neuronal GABA receptors. Trends Neurosci. 20, 578–583.)

p0205

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2000) and Acheta domesticus (Strambi et al., 1998). RDL is very strongly expressed (Harrison et al., 1996), where the pattern of staining is compartmentalized, suggesting that some, but not all, Kenyon cells (the principal interneurons of the mushroom bodies) receive GABAergic inputs (Brotz et al., 1997). The significance of this pattern of expression is discussed in the following section. The distribution of RDL staining corresponds closely to staining for glutamic acid decarboxylase (GAD; the enzyme involved in the synthesis of GABA) (Buchner et al., 1988), GABA, and synaptotagmin (DiAntonio et al., 1993), suggesting that they contribute to the formation of synaptic receptors. Similar patterns of distribution in the mushroom bodies have been observed in the case of the cockroach P. americana (Sattelle et al., 2000) and the cricket A. domesticus (Strambi et al., 1998), where again RDL distribution closely matched GABA staining, although details of the pattern of staining differed somewhat across different species. When expressed in Xenopus oocytes, RDL forms receptors (presumably homomeric, as no endogenous GABAR subunits have been reported in these cells) that respond to GABA with thresholds around 1–10 mM (ffrench-Constant et al., 1993a, 1993b; Hosie et al., 1995a, 1995b) as well as in baculovirus transfected sf9 cells (Lee et al., 1994) and a Drosophila S2 cell line (Millar et al., 1994). These currents reverse close to the equilibrium potential for chloride ions. Responses are dose-dependent and there is little evidence of rapid ( CACA  glycine (Hosie et al., 1995a, 1995b). Similarly, the rank order of potency of agonists acting on RDL stably expressed in an insect cell line is GABA ¼ TACA > CACA  glycine ¼ taurine (Buckingham et al., 1996). The lack of activity of glycine is of interest, since the amino acid sequence of RDL more closely resembles that of glycine receptors than it does of many GABAR subunits (ffrench-Constant and Rocheleau, 1992). RDL is also insensitive to 3-APS yet is activated by isoguvacine (Hosie and Sattelle, 1996a). The high potency of muscimol (Figure 2), the sensitivity of RDL to CACA and isoguvacine, along with its insensitivity to 3-APS, are all features characteristic of insect nervous system GABARs, but not of insect muscle (see above). This supports the observation that RDL is widely expressed in the nervous system but not in muscle. An interesting set of findings resulting from a chance observation of atypical agonist induced currents in RDL led to the discovery of the strong pH sensitivity of RDL. The GABA agonists THIP (4,5,6,7-tetrahydroisoxazolo[5,4-c]pyridin-3-ol) and ZAPA (Z-3-[(aminoiminomethyl)thio]prop-2-enoic acid) appeared to lead to a desensitizing response, followed by a pronounced ‘‘off’’ response upon agonist washout (Matsuda et al., 1996). Further investigation showed this to be the result of acidification of the test saline by the agonists. When this was corrected, more typical responses were observed.

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What is particularly significant about this, however, is that the apparent off response was due to a pH dependency of the efficacy of these two compounds–preacification of the saline greatly reduced the amplitude of the responses to these agonists. The actions of other agonists tested on RDL, including GABA, were pH independent. Other studies had shown that protonation of histidine residues underlies pH dependent actions of GABA on recombinant mammalian receptors (Krishek et al., 1996), as well as the pH dependency of the action of zinc ions (Wang et al., 1995). This points to further differences between the agonist binding sites of RDL and vertebrate GABAA receptors. From the pKa of GABA and THIP at the pH of the uncorrected test solutions (about 4.5), about 76% of the hydroxyls of GABA would be dissociated and about 50% of the hydroxyls of THIP would be dissociated. Thus, some of the differences in pH sensitivity of these drugs may be due to the importance of the dissociated state of the acidic group in ligand binding. Alternatively, these differences may also be attributable to pH dependent states of the histidine residues that form part of the receptor. There are seven histidine residues in the extracellar N-terminal of RDL, and future studies of the effects of point mutations involving each of these residues may help uncover new insights into the agonist actions on RDL. Another key feature in which RDL resembles in situ insect GABARs is its insensitivity to bicuculline. This insensitivity to bicuculline is seen whether RDL is expressed transiently in Xenopus oocytes (Hosie and Sattelle, 1996a) or stably expressed in a Drosophila cell line (Millar et al., 1994). Bicuculline insensitivity is a feature also possessed by the rdl gene cloned from H. virescens (Wolff and Wingate, 1998). Since, as argued above, most insect GABARs are bicuculline insensitive, RDL thus resembles the majority of insect GABARs, including those expressed on cultured Drosophila neurons (Zhang et al., 1994). In some respects, RDL is more similar to GABAC receptors and the recombinant receptors formed by expression of the vertebrate r subunit in their insensitivity to bicuculline and the ability to form functional homo-oligomers. However, it is clear that the agonist site of RDL is dissimilar to that of GABAC receptors in its preferred pharmacophore. At the sequence level, RDL bears no more closer resemblance to GABAC receptors than it does to GABAA receptors, and in fact more closely resembles glycine receptors (ffrench-Constant and Rocheleau, 1992). Thus, while RDL homomers are to be distinguished from GABAA receptors by their bicuculline insensitivity and the low potency of 3-aminosulfonic acid, they are also to be distinguished from GABAC

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receptors by the high efficacy of muscimol and isoguvacine. 5.4.2.4.3.2. The convulsant antagonist site The RDL channel expressed in Xenopus oocytes or the Drosophila S2 cell line is effectively blocked by 1–10 mM concentrations of PTX, TBPS, EBOB, dieldrin, BIDN, and fipronil (ffrench-Constant et al., 1993a, 1993b; Millar et al., 1994; Shirai et al., 1995; Buckingham et al., 1996). For structures of these molecules see Figures 3–5. The IC50 for PTX block of RDL expressed in Drosophila cells was estimated to be 2.5 mM, whereas 10 mM TBPS reduced GABA evoked currents by less than 50%. This higher efficacy of PTX compared to TBPS is also typical of in situ insect GABARs, whereas the reverse is true for vertebrate GABARs. Curiously, the RDL homolog from H. virescens is insensitive to dieldrin (Wolff and Wingate, 1998). The block of RDL mediated GABA responses by PTX has both competitive and noncompetitive features, in that it reduces the maximal response to GABA and shifts the EC50 of the GABA response (Shirai et al., 1995). A similar action of PTX was reported for a crustacean GABAR (Smart and Constanti, 1986). As is the case for insect GABARs in situ, PTX is a more potent blocker of RDL than TBPS (Buckingham et al., 1996), whereas the reverse is true for vertebrate GABAA receptors. The convenience of the availability of a robust, functional homomeric GABAR was exploited by Shirai and coworkers (Shirai et al., 1995) to determine the key structural elements of the PTX molecule that determine its potency. Using the same approach as that on expressed vertebrate GABAA receptors (Anthony et al., 1993) and in situ insect receptors (Anthony et al., 1994), the study agreed with earlier ones in that picrotin and PTX acetate were found to be of less efficacy in blocking the GABA response, whereas fluoropicrotoxin was of similar potency to PTX. Thus, the PTX recognition site of RDL resembles that of in situ insect GABARs. The potential that RDL offers over in situ receptors, namely that the subunit composition is known, could be exploited further in elucidating the mechanisms of action of PTX and related compounds. These structure–function studies did not uncover any significant difference between vertebrate and insect GABARs at the convulsant site, yet such differences presumably exist, as is inferred from the selectivity of certain insecticides known to act at the convulsant site. Such differences did, however, emerge in a subsequent study by Sattelle and colleagues (Hosie et al., 1996), which used a related series of PTX-like compounds, the picrodendrins, to probe the convulsant antagonist site. Picrodendrins

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and related tutin analogs are terpenoid toxins present in extracts of a plant of the Euphorbiaceae, Picrodendron baccatum. These plants are known as ‘‘mata becerro,’’ or calf-killer, in the Dominican Republic where they have been used to kill bed bugs and lice (Ozoe et al., 1994). The picrodendrins are structurally similar to PTX, and share its polycyclic lactone picrotoxane skeleton, acting at a similar site in the channel of vertebrate GABAA receptors (Ozoe et al., 1994). The series of terpenoids studied differed mainly in their substituents at the C9 position. In most cases, these substituents were a spiro-g-butyrolactone ring, providing one of at least two electronegative groups believed to be necessary for picrotoxane activity (Ozoe and Matsumura, 1986). Alteration of these substituents produced differing effects on their potency on RDL when compared to their effectiveness in displacing [35S]TBPS from rat membranes (Ozoe et al., 1994). First of all, most of the terpenoids were significantly less effective at blocking RDL than at displacing TBPS from rat membranes. More importantly, however, there were considerable differences in their rank order of potency on RDL compared to rat GABARs. Molecules that were most potent against RDL shared an olefin binding that brought the side chains closer to the plane of the spiro-g-butyrolactone ring than the other compounds. This study succeeded in showing the importance of the position of the electronegative centers in the PTX-like family, which had not emerged from studies restricted to the C4 substituents and the isopropenyl moiety. The bicyclic dinitrile BIDN, developed in the face of a need for a new, specific ligand for the insect convulsant site, is a potent blocker of recombinant RDL GABA receptors. At 1 mM, BIDN reversibly reduces the amplitude of RDL mediated GABA responses in Xenopus oocytes, with only a slight dependence on membrane potential (Hosie et al., 1995b). This action was dose dependent, with an IC50 that depended upon the concentration of GABA used. When tested against 20 mM GABA the IC50 for BIDN action was estimated to be 1.0 mM, whereas when tested against 100 mM GABA it was estimated at 20 mM. This is suggestive of allosteric interactions between the BIDN and GABA ligand binding sites. Like PTX, the action of BIDN displayed both competitive and noncompetitive elements, in that it reduced the maximum GABA induced currents and shifted the dose–response curve to the right, pointing to allosteric actions of BIDN. BIDN also reversibly blocks RDL mediated responses in the S2-RDL and S2-RDL(A302S) cell lines (Buckingham et al., 1996). In both the

Xenopus oocyte and S2 cell line expression systems, it was noticed that GABA responses in the presence of BIDN lacked the desensitization that was so clearly present in control responses (Buckingham et al., 1996). One plausible interpretation of this might simply be that, like PTX, BIDN exerts its effects through an enhancement of desensitization, but favoring a different state than PTX. Evidence that BIDN and fipronil both operate by reducing open-state probability, but through different mechanisms, also comes from single channel studies (Grolleau and Sattelle, 2000). RDL expressed in Xenopus oocytes and in a Drosophila cell line is sensitive to block by the phenylpyrazole insecticide, fipronil (Millar et al., 1994; Hosie et al., 1995a). In Xenopus oocytes, RDL mediated responses to 50 mM GABA are reduced by up to 80% by 100 mM fipronil. Recovery from fipronil block was very slow, and still incomplete after 10 min washout. This differs from the actions of both PTX and BIDN, which reverse completely and rapidly. Like the block by BIDN (Hosie et al., 1995b), fipronil block was voltage independent (Hosie et al., 1995a). A further similarity to the actions of both BIDN and PTX is that the action of fipronil consists of both competitive and noncompetitive elements, and is also dependent on the concentration of agonist. Thus, fipronil, BIDN, and PTX share many features (voltage independence, agonist dependence, and mixed mode of action) suggesting that they operate through similar mechanisms. However, it must be noted that whereas PTX and the cyclodienes are competitive inhibitors of [3H]EBOB binding, fipronil is a noncompetitive displacer of [3H]EBOB binding, at least in M. domestica (Deng et al., 1991, 1993). The possible significance of this in understanding the mechanisms of insecticide resistance is discussed below. Differences in the action of fipronil and BIDN on RDL were examined in a single channel patchclamp study (Grolleau and Sattelle, 2000). The ability of RDL to form functional homomers simplifies the interpretation of experiments on the effects of the A302S mutation on single channel properties of the channel. When applied to outside-out patches from S2-RDL cells, GABA (50 mM) evoked inward currents that were completely blocked by 100 mM PTX. As had been reported for outside-out patches of Xenopus oocytes expressing RDL, the current– voltage relationship showed slight inward rectification at positive potentials. The conductance of the single channel currents was around 36 pS, but interestingly a 71 pS conductance was observed when the application of GABA was longer than 80 ms. When either 1 mM fipronil or 1 mM BIDN was present in

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the external saline, all the GABA gated channels were completely blocked. At lower concentrations, near their respective IC50 values, the duration of channel openings was shortened. Interestingly, the blocking action of BIDN resulted in the appearance of a novel channel conductance (17 pS). Examination of the effects of coapplication of these two convulsants (applying 300 nM BIDN with fipronil concentrations ranging from 100 to 1000 nM) revealed an additional BIDN induced dose dependent reduction in the maximum open probability. Thus, while both BIDN and fipronil shorten the duration of wild-type RDLac GABAR channel openings, they appear to do so by acting at distinct sites. The findings reported to date on the actions of convulsants on RDL agree very closely with their actions on in situ insect GABARs. The agonist dependence and mixed competitive/noncompetitive action are consistent with a model of PTX action in which the PTX molecule binds preferentially to an agonist-bound conformational state of the receptor and hence increases the probability of a transition into a desensitized state. Such a conclusion was advanced to explain similar properties of PTX action at the crayfish neuromuscular junctional GABARs (Smart and Constanti, 1986) and the finding that the rate of development of a steadystate blockade of GABARs on rat sympathetic neurons was enhanced when GABA is applied in brief, rapidly repeated doses (Newland and Cull-Candy, 1992). RDL has also been used to predict cross-resistance to insecticides acting at the convulsant site. Dieldrin resistant RDL mutant GABARs are not only comparatively insensitive to PTX, but are also insensitive to BIDN (Hosie et al., 1995b). Similar results were obtained for fipronil (Hosie et al., 1995a) although other workers reported different results (Wolff and Wingate, 1998). Furthermore, a novel tricyclic dinitrile, KN244, blocks RDL and the sensitivity of dieldrin resistant RDL to it is reduced 100-fold (Matsuda et al., 1999). 5.4.2.4.3.3. Allosteric modulators Responses mediated by RDL are sensitive to potentiation by benzodiazepines, but their benzodiazepine pharmacology differs from in situ insect receptors. Insect GABA responses are potentiated by Ro 5-4864 and flunitrazepam. Responses to GABA mediated by RDL expressed in Xenopus oocytes (Hosie and Sattelle, 1996b) were potentiated by micromolar concentrations of Ro 5-4864. In contrast, the same receptor expressed in the Drosophila S2 cell line appeared to be insensitive to Ro 5-4864 (Millar

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et al., 1994), although enhancement was obtained when the concentration of the benzodiazepine was increased to 100 mM (Buckingham et al., 1996b). This illustrates that caution must be exercised in interpreting negative data until a wide range of ligands is tested. However, flunitrazepam, which is active at GABARs on cockroach motor neuron, Df, at micromolar concentrations, was found to be ineffective on RDL in either expression system even at concentrations of 100 mM (Millar et al., 1994; Hosie and Sattelle, 1996b). This represents a departure from most in situ insect receptors studied to date. Significantly, the identified motor neuron Df of the cockroach stains positively to the RDL antibody (Sattelle, personal communication), suggesting that in this cell at least, RDL may coassemble with another subunit. Flunitrazepam is also reported to have no effect on a GABAR cloned from H. virescens (Wolff and Wingate, 1998). Nonetheless, the comparatively low sensitivity of RDL to benzodiazepines, coupled with its sensitivity to the ‘‘peripheral’’ benzodiazepine, Ro 5-4864, are both characteristic of insect receptors compared to that of vertebrates. That RDL should differ from in situ receptors principally at the benzodiazepine site is not surprising, given the wide variation in benzodiazepine pharmacology of GABARs generally, and may indicate that RDL coassembles with another subunit subtype in situ. Since only two other GABARs have been cloned from Drosophila to date, these are the obvious candidates for the other subunit. However, it is unlikely that LCCH3 is the additional subunit, since, as described above, receptors formed by coexpression of RDL with LCCH3 are quite unlike insect receptors. In any case, the lack of overlap of expression of these two subunits at any stage of development makes it highly unlikely that they coassemble in vivo. The question of whether RDL forms a homomer in situ is discussed in a separate section below. Unfortunately, coexpression of RDL with GRD has yet to be described. While expression studies on the agonist pharmacology of three of the four RDL splice variants has been described in some detail (Chen et al., 1994; Hosie and Sattelle, 1996a), benzodiazepine pharmacology of these variants, or combinations of them, has not yet been assayed. Like in situ insect receptors, RDL is potentiated by barbiturates. In both the Xenopus oocyte and the Drosophila S2 cell line expression systems, RDL mediated GABA responses were enhanced by either sodium phenobarbitone or sodium pentobarbitone by up to 50% of control values (Hosie and Sattelle, 1996b; Buckingham et al., 1996). Similar results were reported for the RDLbd splice variant, whose responses were enhanced up to fivefold (Belelli et al.,

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1996a). Similarly, the RDL equivalent cloned from H. virescens is also subject to barbiturate (phenobarbital) enhancement (Wolff and Wingate, 1998). In contrast to vertebrate GABARs, which are also sensitive to barbiturates, elevation of the barbiturate to high (1 mm) concentrations did not evoke Drosophila RDL mediated currents in the absence of GABA (Belelli et al., 1996a; Buckingham et al., 1996). It is therefore likely that the two components of barbiturate action on vertebrate receptors – the enhancement of the GABA response and the direct opening of the channel in the absence of GABA – are mediated by two separate sites, of which only one may be present on insect GABARs. RDL is comparatively insensitive to steroids. The steroid, 5a-pregnan-3a-ol-20-one, enhances human a3b1g2L receptors, but at high concentrations had only a minimal effect on RDLbd (Belelli et al., 1996) and RDLac (Millar et al., 1994; Hosie and Sattelle, 1996b). Insect GABARs in situ are also highly insensitive to steroids (Rauh et al., 1990). The anticonvulsant, loreclezole, is a potentiator of vertebrate recombinant GABARs containing the b2 or b3 subunit (Wafford et al., 1995), but not of those containing a b1 subunit (Wingrove et al., 1994). This is attributed to the presence of a serine in the b2 and b3 subunits, which is an asparagine at the equivalent position in the b1. Curiously, although a mutation of this serine to a methionine abolishes loreclezole sensitivity (Stevenson et al., 1995), RDL (which has methionine at this position) is sensitive to loreclezole (Belelli et al., 1995; Hosie and Sattelle, 1996b). RDL is also sensitive to a number of other allosteric modulators. Isoflurane enhances GABA responses mediated by expressed RDL and blocks at higher concentrations. This block is not seen for the cyclodiene resistant A302S mutant, suggesting that the block is due to an interaction at the PTX site (Edwards and Lees, 1997). These data amply illustrate the usefulness of the cloned RDL receptor in advancing our understanding of insect GABARs. More than this, however, RDL, because of its ability to form functional homomers, has also been of use in structure/function studies on allosteric modulator sites of vertebrate GABARs. For instance, RDL is insensitive to the anesthetic etomidate, as are vertebrate b1 subunits. Vertebrate b2 and b3 subunits, however, do confer sensitivity to etomidate and this is attributable to the presence of an asparagine residue at equivalent positions in a transmembrane-spanning domain, which is a methionine at the equivalent position in RDL. Mutating M314 to an asparagine in RDL rendered the expressed receptors sensitive to

enhancement by etomidate (McGurk et al., 1998). As in vertebrate recombinant receptors, this sensitivity is stereospecific, whereas the native sensitivity to pregnane steroids or pentobarbitone remained unchanged. Conversely, the N289M mutation introduced into vertebrate b2 abolished the etomidate sensitivity. Such complementarity of effects of mutation encourages confidence in interpreting experiments using an invertebrate receptor as a model of vertebrate receptors, especially in view of the comparative simplicity of interpreting data obtained using homomers. 5.4.2.4.3.4. Other (RDL) ligands RDL, like GABARs of the motor neuron, Df, of the cockroach (Bai, 1994), is insensitive to zinc (Hosie and Sattelle, 1996b), which blocks many vertebrate GABAA receptors (Hosie et al., 2003), as well as r receptors. 5.4.2.4.4. Conclusions There can be little doubt that the availability of RDL has accelerated the pace of research into the molecular pharmacology of insect GABARs. The chief advantage that it confers, in addition to the convenience of performing experiments on oocytes, is that hitherto studies aimed at uncovering the mechanisms of drug–receptor interaction for insect GABARs have been hindered by the fact that in no case is the subunit composition of any in situ GABAR known. It is of great significance that a number of different laboratories have confirmed independently that whatever the expression system chosen (Xenopus oocytes, Drosophila S2 cells, Spodoptera Sf9 cells), the pharmacology of the expressed RDL receptor does not differ significantly. Indeed one paper (Buckingham et al., 1996) examined this issue directly. This encourages confidence that RDL is not coexpressing with GABAR subunits native to the respective expression systems, and that we can be reasonably certain that the subunit composition of the expressed RDL receptor is a homomeric pentamer. To appreciate the significance of such an advantage, we have only to look at the advances in understanding of nicotinic receptors that emerged from research using another homomer forming receptor, the vertebrate nicotinic a7 receptor. Since RDL homomers so closely mimic in situ GABARs, and since RDL antibody staining corresponds closely to GABA immunoreactivity, it is likely that RDL contributes to most of the GABARs in the insect brain. In view of the conclusion drawn above that there is pharmacological diversity of subtypes of GABARs in insects, the question of the source of this diversity remains to be addressed. The occurrence of alternative splicing and RNA

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editing may add to functional diversity of the small Drosophila GABAR gene family. Evidence for this possibility is discussed in the following section. 5.4.2.4.5. Alternative splicing in GABA receptor subunits RDL is expressed in the form of four splice variants, arising from alternative splicing of two of its nine exons (exons 3 and 6). These four distinct polypeptides show characteristic structural features of a GABAR subunit. The regions in which the splice variants differ align closely with regions known to be determinants of agonist potency in vertebrate ionotropic receptor subunits. This is highly unusual. Although there are many examples of alternatively spliced GABAR subunit genes in vertebrates, the splicing usually occurs in their large intracellular loops. All four of these splice variants are expressed in embryonic Drosophila (ffrench-Constant and Rocheleau, 1992), yet their functional roles remain to be determined. Since the pharmacological diversity observed among vertebrate GABAA receptors arises at least in part from the assorted assembly from different subunit isoforms, the alternate splicing of the rdl gene may serve a physiological role by increasing receptor diversity. Since RDL appears to be a component of the majority of insect GABARs, such a splicing mechanism may represent another route to subunit diversity in a small GABAR gene family. Evidence that alternative splicing confers pharmacological diversity comes from expression studies that have described pharmacological differences between these variants at the agonist site. For example, the three splice variants tested to date in functional expression studies (RDLac, RDLad, RDLbd) all differ in their EC50 values for GABA (Hosie and Sattelle, 1996a). The region in which these variants differ include loop F of the GABA binding site. 5.4.2.4.6. Does RDL form a homomer in situ? Despite our limited knowledge of the pharmacology of native GABARs of Drosophila, it is clear that RDL homomers mimic closely the pharmacology of insect GABARs. The general features that distinguish most insect GABARs from those of vertebrates – insensitivity to bicuculline, comparative insensitivity to benzodiazepines, sensitivity to the peripheral benzodiazepine Ro 5-4864, insensitivity to TBPS, and sensitivity to isoguvacine – are all seen in RDL homomers. The only challenge to this, the insensitivity of RDL to flunitrazepam, is perhaps not particularly serious in view of the diversity of benzodiazepine pharmacology amongst GABAA receptors studied to date and the acute dependence of benzodiazepine pharmacology upon

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subunit composition. Where this difference in benzodiazepine pharmacology gains significance, though, lies in the observation that a Drosophila anti-RDL antibody stains motor neuron Df of the cockroach (Sattelle, personal observation), which is known to possess GABARs sensitive to micromolar concentrations of flunitrazepam (Sattelle et al., 1988). Since RDL, which is flunitrazepam insensitive, is expressed on Df, which is flunitrazepam sensitive, this difference hints at the existence of another subunit subtype in these cells. Single channel recordings also provide reason to suspect that RDL may not always exist as a homomer in situ. Single, GABA gated channels recorded from Xenopus oocytes injected with rdl cRNA were inwardly rectifying, with inward and outward currents of