Advances in Agrochemicals: Ion Channels and G Protein-Coupled Receptors (GPCRs) As Targets for Pest Control [2] 0841232601, 9780841232600

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Advances in Agrochemicals: Ion Channels and G ProteinCoupled Receptors (GPCRs) as Targets for Pest Control Volume 2: GPCRs and Ion Channels

ACS SYMPOSIUM SERIES 1265

Advances in Agrochemicals: Ion Channels and G ProteinCoupled Receptors (GPCRs) as Targets for Pest Control Volume 2: GPCRs and Ion Channels Aaron D. Gross, Editor Virginia Polytechnic Institute and State University Blacksburg, Virginia, United States

Yoshihisa Ozoe, Editor Shimane University Matsue, Shimane, Japan

Joel R. Coats, Editor Iowa State University Ames, Iowa, United States Sponsored by the ACS Division of Agrochemicals

American Chemical Society, Washington, DC Distributed in print by Oxford University Press

Library of Congress Cataloging-in-Publication Data Names: Gross, Aaron (Aaron D.), editor. | Ozoe, Yoshihisa, editor. | Coats, Joel R., 1948- editor. | American Chemical Society. Division of Agrochemicals. Title: Advances in agrochemicals : ion channels and G protein-coupled receptors (GPCRs) as targets for pest control / Aaron D. Gross, editor (Virginia Polytechnic Institute and State University, Blacksburg, Virginia, United States), Yoshihisa Ozoe, editor (Shimane University, Matsue, Shimane, Japan), Joel R. Coats, editor (Iowa State University, Ames, Iowa, United States) ; sponsored by the ACS Division of Agrochemicals. Description: Washington, DC : American Chemical Society, [2017]- | Series: ACS symposium series ; 1264, 1265 | Includes bibliographical references and index. Identifiers: LCCN 2017049010 (print) | LCCN 2017049810 (ebook) | ISBN 9780841232587 (ebook) | ISBN 9780841232570 (v. 1) | ISBN 9780841232600 (v. 2) Subjects: LCSH: Agricultural pests--Control. | Agricultural chemicals. | Ion channels. Classification: LCC SB950 (ebook) | LCC SB950 .A35 2017 (print) | DDC 628.9/6--dc23 LC record available at https://lccn.loc.gov/2017049010

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Foreword The ACS Symposium Series was first published in 1974 to provide a mechanism for publishing symposia quickly in book form. The purpose of the series is to publish timely, comprehensive books developed from the ACS sponsored symposia based on current scientific research. Occasionally, books are developed from symposia sponsored by other organizations when the topic is of keen interest to the chemistry audience. Before agreeing to publish a book, the proposed table of contents is reviewed for appropriate and comprehensive coverage and for interest to the audience. Some papers may be excluded to better focus the book; others may be added to provide comprehensiveness. When appropriate, overview or introductory chapters are added. Drafts of chapters are peer-reviewed prior to final acceptance or rejection, and manuscripts are prepared in camera-ready format. As a rule, only original research papers and original review papers are included in the volumes. Verbatim reproductions of previous published papers are not accepted.

ACS Books Department

Contents Preface .............................................................................................................................. ix 1.

Variations in the Insect GABA Receptor, RDL, and Their Impact on Receptor Pharmacology .......................................................................................... 1 Jennina Taylor-Wells and Andrew K. Jones

2.

Insecticide Resistance in Rice Planthoppers ........................................................ 23 Toshifumi Nakao

3.

The GABA Antagonist γ-Benzene Hexachloride and its Polychlorinated Cyclohexane Analogs ............................................................................................. 41 Keiji Tanaka, Takaaki Sakamoto, Takaaki Iwai, Kou Kuroda, Karin Nagasaki, Yoshihisa Ozoe, Miki Akamatsu, and Kazuhiko Matsuda

4.

Potential of GPCR-Targeting Insecticides for Control of Arthropod Vectors ..................................................................................................................... 55 Shruti Sharan and Catherine A. Hill

5.

Subunit-Specific Modulatory Functions Are Conserved in an Interspecies Insect GABAB Receptor Heteromer ..................................................................... 85 S. Blankenburg, S. Balfanz, A. Baumann, and W. Blenau

6.

Functional Characterization of Dopamine and Neuropeptide G Protein-Coupled Receptors from the Silkworm Bombyx mori By Aequorin Bioluminescence-Based Calcium Assay ............................................................. 109 Hiroto Ohta, Kanako Mitsumasu, Toshinobu Yaginuma, Yoshiaki Tanaka, and Kiyoshi Asaoka

7.

Molecular Pharmacology and Physiology of Insect Biogenic Amine Receptors ............................................................................................................... 127 Jia Huang

8.

A Review of Aminothiazoline Chemistry ........................................................... 139 Barbara Wedel, Wolfgang von Deyn, Sebastian Soergel, Matthias Pohlman, Douglas Anspaugh, Ramani Kandasamy, Fae Malone, Daniel Houtz, Nancy Rankl, John Dorsch, Lynn Stam, Brecht London, Ronan le Vezouet, Christopher Koradin, and Markus Kordes

Editors’ Biographies .................................................................................................... 149

vii

Indexes Author Index ................................................................................................................ 153 Subject Index ................................................................................................................ 155

viii

Preface Pest management often depends on the use of chemical insecticides that affect important physiological and pharmacological processes in arthropods. The development of chemical insecticides has primarily attacked the arthropod nervous system by targeting and disrupting the function of ligand-gated or voltage-gated ion channels. The use of broad classes of synthetic chemistries with common modes of action, along with the widespread, and sometimes improper, use of chemical insecticides has resulted in the urgent need to find new chemistry and novel or underutilized targets, such as G-protein-coupled receptors (GPCRs) for screening, discovery, and development of novel agrochemicals. In addition, we are presenting several recent updates to our knowledge of vital ion channels. Through our authors’ highlighting of rapidly expanding research on GPCRs, as well as more traditional targets, we hope to spur new lines of research and design of new synthetic or biorational agrochemicals. We offer here chapters that describe investigations on GPCR’s that document the substantial progress that has been made recently in understanding the mechanisms of ligand interactions at the receptors, the downstream cascades, and the exciting promise that this large class of receptors has as targets for new agrochemical products. Multiple experts focus on the latest developments on characterization of their molecular pharmacology, on dopamine-specific GPCRs, and on the breadth of various GPCRs as future targets. Our authors have also contributed multiple chapters on insect GABA receptors, addressing their characterization, pharmacology and specificity, as well as specific insecticides and analogs that are active at those receptors. There is also a review of the aminothiazolines as agrochemicals, and a chapter focused on the ever-growing problems from insecticide-resistant pests, in this case, specifically in rice plant hoppers. As the challenges of controlling arthropod pests loom larger each year, we hope this book will serve to ignite new directions, new collaborations, and novel solutions for the security of our food and fiber, as well as the protection of public health. ADG dedicates this book to his family and his great academic mentors. YO dedicates this book to the late Professors M. Eto and F. Matsumura, and his family. JRC dedicates this book to his grandchildren.

ix

Aaron D. Gross Virginia Polytechnic Institute and State University Department of Entomology 207 Latham Hall (MC 0390) 220 Ag Quad Lane Blacksburg, Virginia 24061, United States [email protected] (e-mail)

Yoshihisa Ozoe Faculty of Life and Environmental Science Shimane University Matsue, Shimane 690-8504, Japan [email protected] (e-mail)

Joel R. Coats Distinguished Professor of Entomology and Toxicology Department of Entomology 116 Insectary Iowa State University Ames, Iowa 50011-3140, United States [email protected] (e-mail)

x

Chapter 1

Variations in the Insect GABA Receptor, RDL, and Their Impact on Receptor Pharmacology Jennina Taylor-Wells and Andrew K. Jones* Faculty of Health and Life Sciences, Department of Biological and Medical Sciences, Oxford Brookes University, Headington, Oxford OX3 8NZ, United Kingdom *E-mail: [email protected].

The resistance to dieldrin (RDL) receptor is an insect γ-aminobutyric acid (GABA) receptor, characterized by the dieldrin resistance mutation that was pivotal to understanding target based insecticide resistance. RDL is the target for various non-competitive antagonists, including dieldrin and fipronil, as well as novel acting compounds such as the meta-diamides and isoxazolines. Therefore the RDL receptor has returned to center stage as a relevant and effective insecticide target. Our understanding of the function of RDL in vivo is still unfolding, with the discovery of species specific post-transcriptional modifications such as alternative splicing and RNA editing, modifications shown to influence the pharmacology of the receptor. Exposing these receptors to insecticides also evokes ever evolving mechanisms of mutagenesis, and a number of contributory mutations have been identified both in field and laboratory resistant insects, occurring in parallel to the dieldrin resistance mutation. We present an overview of these variations and discuss the impact on the pharmacology of GABA and various insecticides.

© 2017 American Chemical Society

Introduction The targeting of insect neuronal receptors is one of the main mechanisms of insecticidal action. However, our understanding of these neuronal receptors and their functions in vivo are still unfolding, with the continuing discovery and functional analysis of post-transcriptional modifications such as alternative splicing and RNA editing. By the same token, exposing these receptors to insecticides evokes ever evolving mechanisms of mutagenesis by nature, contributing to insecticide resistance or offsetting the fitness costs of other mutations. An overview of these variations and their effects on receptor pharmacology are provided on the resistance to dieldrin (RDL) receptor, the most studied of the insect γ-aminobutyric acid (GABA) receptors and the target for both historic and novel non-competitive antagonists (NCAs) (1). The RDL receptor is involved in rapid inhibitory synaptic transmission, utilizing GABA as its neurotransmitter. As the first invertebrate GABA receptor to be identified, most sequence and functional analysis has been conducted in the fruit fly model Drosophila melanogaster, in which the subunit was first identified and functionally expressed (2, 3). The RDL receptor plays a key role in various processes, most notably the regulation of sleep (4), aggression (5) and olfaction (6, 7), and in D. melanogaster, RDL is expressed both in the embryonic and adult central nervous system (8, 9). As a member of the cys-loop ligand-gated ion channel (cysLGIC) superfamily, the RDL receptor consists of a pentameric subunit structure, centered around a central pore. In turn, the RDL subunit is composed of extracellular N- and C-termini, four transmembrane domains (M1 – M4), the second of which lines the ion channel; and a large intracellular loop between M3 and M4 (10). The agonist binding site is located in the N-terminal extracellular domain and consists of distinct regions (loops A-F) (11). Also in the N-terminal domain is the dicysteine loop, which is characteristic of the cysLGIC superfamily. In addition to RDL, the cysLGIC superfamily contains other receptor targets for insecticides, including the glutamate gated chloride channels and nicotinic acetylcholine receptors (1, 12, 13). GABA binds at the subunit interface, altering the conformation of the receptor and allowing the passage of chloride ions, which initiate the cascade of inhibitory action. By this function, the insect RDL receptor is most related to vertebrate GABAA receptors, though pharmacologically it does have some differences (14, 15). The RDL receptor was identified and characterized following the discovery of a mutation located in the ion channel pore forming M2 (amino acid position 301) of D. melanogaster Rdl. This mutation rendered the RDL receptor resistant to the antagonistic effects of 10 μM dieldrin, which is a cyclodiene insecticide (3). The Rdl subunit has since been cloned from several insect orders, including those of agricultural pests (e.g. red flour beetle Tribolium castaneum and planthopper Laodelphax striatellus (16, 17)), pests afflicting domesticated animals (cat flea, Ctenocephalides felis (18)), human and animal disease vectors (house fly Musca domestica, and mosquitoes Anopheles gambiae and Aedes aegypti (19–21)) and beneficial species (miridbug Cyrtorhinus lividipennis and honeybee Apis mellifera 2

(22, 23)). In most cases, an insect has only one Rdl gene, however, there are exceptions, such as the presence of two Rdl genes in the pea aphid (Acyrthosiphon pisum) (24) and the diamondback moth (Plutella xylostella) (25), while the silk worm (Bombyx mori) possesses three Rdl genes (26). The coding sequence of Rdl is remarkably conserved between diverse insect species, usually showing 70-90% identity at the amino acid level (27, 28). The M2 mutation in the RDL subunit (commonly referred to as the A2′ mutation, which signifies an alteration in the second amino acid position of the ion channel domain) (15) has been extensively studied since its discovery, and is utilized as a diagnostic marker for resistance (29). However, several other RDL subunit mutations have recently been discovered in field and laboratory selected resistant insects, and interestingly always in parallel with the A2′ mutation (30–32). The RDL receptor is a target for a number of neuromodulatory and insecticidal compounds including lindane (33), picrotoxin (34), cyclodienes (e.g. dieldrin, endosulfan) (2), phenylpyrazoles (e.g. fipronil) (35) and macrocyclic lactones (e.g. ivermectin) (36). Recently, the RDL receptor has returned to center stage as a novel target for the isoxazolines (37, 38), meta-diamides (39) and meroterpenoid chrodrimanins (40), that target different binding sites and have the distinct advantage of acting on RDL bearing subunits containing the M2 resistance mutation (15, 40). The study of RDL receptor is facilitated by the ability to express high levels of the homomeric receptor in Xenopus laevis oocytes and other expression systems (41–44), but the exact composition of the subunits in vivo remains unknown. These homomeric receptors maintain most of the pharmacological properties of native insect GABA receptors but their response to some benzodiazepines is different (14, 15), suggesting that other subunits may co-assemble with RDL in vivo. This is supported by in vitro studies showing that RDL can co-express with the GABA receptor subunits GRD (GABA/glycine-like receptor of Drosophila) (45) and LCCH3 (ligand-gated chloride channel homologue 3) (46) as well as the glutamate gated chloride channels (GluCls) (21, 47). However, there is other evidence to suggest that these subunits may not necessarily co-assemble in vivo (48, 49). The RDL receptor is likely to be far more complex than is suggested by the common perception that it is a homomeric. This is because alternative splicing and RNA editing have been found to increase the diversity of the Rdl transcriptome (15, 50). In addition, evidence has come to light that these variations can affect the pharmacology of the receptor and influence the actions of insecticides (51, 52). In the following chapter we will consider the recent molecular variations identified in the insect RDL receptor and discuss the impact of these variations on the receptor pharmacology of GABA and various insecticides.

3

Environmentally-Induced Variations: The Effects of RDL Resistance Mutations on Receptor Pharmacology It is 24 years since the original dieldrin resistance mutation at position 301 (A2′S/G) was identified in an RDL subunit (3), and it is one of the most significant examples of direct target site resistance to date. Dieldrin was removed from the market several decades ago, but may persist in the environment (53). This combined with the potential for cross-resistance with currently used insecticides such as endosulfan and fipronil (32, 54), may suggest why the dieldrin resistance mutation still persists in insect populations (55, 56). With novel insecticides emerging that target the RDL receptor, albeit with different mechanisms of action (57), it seems timely to review the presence of novel insecticide associated mutations that have been discovered in recent years (summarized in Table 1). The amino acid alanine in the channel forming M2 domain, corresponding to position 301 (A2′) in the D. melanogaster RDL subunit, is mutated most often to serine (18, 58, 59) or glycine (58, 60) to confer resistance to channel blocking insecticides such as dieldrin and at varying levels to fipronil (15, 58, 61). It was recently found that the glycine mutation in fact facilitated a greater resistance to fipronil than the serine mutation (both in vivo and in vitro) in D. melanogaster (62). More recently, in 2010 and 2011, another mutation at the same site was observed in two fipronil resistant field populations of planthopper, in which alanine at the same position was mutated to asparagine (A2′N) (44, 63). The first mutation reported to co-exist with the A2′ mutation was T350M (Table 1), isolated from a laboratory selected Drosophila simulans population resistant to dieldrin, also showing a high level of resistance to fipronil (31). This mutation, located in the M3 domain, was always found in the presence of A2′G. Functional experiments with electrophysiology applied to expressed RDL subunit isoforms in X. laevis oocytes revealed that the T350M mutation contributed to fipronil resistance, as the IC50 of the A2′G/T350M isoform for fipronil (221 nM) was significantly increased from that of A2′G alone (93 nM) and wild-type (31 nM) (31). Furthermore, expression of an RDL subunit isoform containing only T350M also showed reduced sensitivity to fipronil (IC50 215 nM), suggesting that the mutation may also individually contribute to resistance. The Drosophila T350 double resistance profile was also found to be conserved in RDL subunits of the malaria mosquito An. gambiae, that were highly resistant to dieldrin (20). Here, the M3 mutation T345M in the An. gambiae RDL subunit (Table 1), which is equivalent to T350M in D. simulans, was also found in the presence of the A2′G mutation (20). As the mosquitoes in this study were collected from wild populations showing phenotypic resistance to dieldrin, the mutations observed here may represent resistance mutations present in the field. In contrast with the double mutant RDL isoform from D. simulans however, the T345M mutation in An. gambiae did not appear to contribute to fipronil resistance, as a double mutation or individually (20). This supports observations in transgenic D. melanogaster, whereby fipronil resistance was not heightened by the presence of the T-M mutation, both alone and in combination with A2′G (62). It is speculated that the T-M mutation in An. gambiae may therefore play a structural rather than functional role, potentially offsetting the 4

fitness costs imposed by A2′G (20, 62). Modeling of the receptor indicates that the two mutations are indeed in close proximity and hence capable of functional interaction (62). The EC50 of A2′G (60 μM) for GABA was also found to be significantly lower than that of the A2′G/T345M double mutant (198 μM), indicating that the T-M mutation may serve to offset the heightened sensitivity to GABA, that would otherwise be detrimental to neuronal signaling (20). A D. melanogaster Rdl subunit mutation, M360I, in the intracellular loop between M3 and M4, was observed alongside the A2′S mutation (Table 1) in a gene duplication that contained one wild type copy and one double mutant (A2′S/ M360I) copy of Rdl (64). This duplication was associated with intermediate levels of dieldrin resistance and heat shock recovery, suggesting that the duplication may offset the temperature sensitive fitness cost associated with the A2′S mutation (64). Interestingly, M360 can be recoded to valine through RNA A-to-I editing (51). It was found that in the duplicated Drosophila Rdl, there were greater levels of RNA editing at I360 than at the wild-type M360 copy (64), which may suggest a link between RNA editing and the A2′S resistance mutation. However, the individual contribution of M360I to insecticide resistance has yet to be determined at a molecular/functional level. In Anopheles funestus Rdl, a V327I substitution was found to be associated with the A2′S mutation in field collected samples of dieldrin resistant mosquitoes in Cameroon and Burkina Faso (Table 1) (56). The valine to isoleucine substitution is present in the loop between the M2 and M3 domains. Interestingly, the mutation was always found in conjunction with the A2′S mutation, but at a lower frequency than was observed for A2′S (56). The link between this mutated isoform and the resistance phenotype, as well as the functional consequences of this mutation, have however not as yet been investigated. A number of different contributory mutations to the less common A2′N mutation have also been identified Rdl subunits from various species of planthopper (Table 1) (65–67). An R340Q mutation was found in the presence of A2′N in an RDL subunit of fipronil resistant Sogatella furcifera collected from a rice paddy in Japan (65). The mutation is present in the intracellular loop between the M3 and M4 domains, in relative close proximity to the T345M/T350M mutations found in An. gambiae and D. simulans (20, 31). As with the V327I mutation in An. funestus RDL, the R340Q mutation was present at a lower frequency than A2′N, being present in only 9 of 17 cDNA clones carrying the A2′N mutation (65). Using a membrane potential assay and Drosophila Mel-2 cells stably expressing the homomeric RDL mutant isoforms, it was shown that the R340Q did not contribute significantly to fipronil resistance, both in combination with A2′N and alone. This result, combined with the location in the intracellular loop suggest, as with the T345M/T350M mutation, that this substitution may have a more subtle structural role and/or play a role in offsetting the fitness costs of the A2′N mutation. In the brown planthopper Nilaparvata lugens, another recent study revealed a mutation in combination with A2′S, a Q359E mutation between M3 and M4 of the RDL subunit (Table 1) (30). The mutations were observed when field collected insects were driven to resistance using the fiprole insecticide ethiprole. Ethiprole has particular relevance to N. lugens, as along with fipronil it has 5

replaced imidacloprid as a method of insect control in Asia, following resistance to neonicotinoids (68). There were differential effects of the A2′S mutation on the sensitivity of ethiprole and fipronil, a somewhat unexpected finding as the structures differ only by an ethylsulfinyl substituent in ethiprole, compared to a trifluoromethylsulfinyl moiety in fipronil (for structures see reference (69)). Both in vitro (homomeric RDL expressed in X. laevis oocytes) and in vivo (D. melanogaster insecticide bioassays) methods demonstrated that the A2′S mutation contributes to ethiprole resistance (30). This effect was most pronounced in vivo, whereby D. melanogaster with the A2′S mutation exhibited 4000-fold resistance in comparison to the wild-type strain. The Q359E mutation, however, did not contribute significantly to resistance in vitro, indicating that the A2′S mutation is the main mechanism of target site resistance. The authors speculate that the presence of the Q359E mutation may be owing to a structural linkage between the two mutations, or that the Q359E mutation confers a fitness advantage. It is difficult to envisage how Q359E may offset any fitness associated with A2′S considering that the mutation, both singly and as a double mutant, did not impact on the potency of GABA (30). Secondary mutations were also found in the M2 domain of the RDL subunit from the planthopper L. striatellus (66). Sequencing of the Rdl gene from field collected insects driven to 87-fold resistance to fipronil revealed the A2′N mutation in combination with either an R305Q or R305W mutation (Table 1). The strain exhibited low cross-resistance to dieldrin and endosulfan and resistance was not affected by detoxification enzymes, suggesting that target site mutations are likely to be responsible for fipronil resistance (66). In N. lugens, an R299Q mutation, also in the M2 domain of the RDL subunit, was recently identified in addition to the A2′S mutation (Table 1) (67). The A2′S/R299Q double mutant was found in insect populations from China, Thailand and Vietnam, with R299Q occurring at a lower frequency than A2′S. In addition, the R299Q mutation was identified during laboratory selection with fipronil (up to a resistance ratio of 237-fold) after the A2′S mutation had reached 100% penetrance. This suggests that R299Q is a secondary mutation potentially associated with persistent fipronil exposure. The mutations, as in previous studies, were only found in parallel, which the authors speculate may be because of the fitness costs associated with the R299Q mutation alone. Indeed, expression of the R299Q mutant subunit expressed as a homomeric receptor individually reduced the GABA potency almost 11 times (EC50 = 413 μM) to that of wild type receptors (38 μM), suggesting a high fitness cost. This is a much larger reduction in potency than was observed with the A2′S mutation (EC50 = 19 μM, 2 times lower). In contrast, the double mutant receptor restored the GABA potency to comparable levels with the wild-type (EC50 = 54 μM), perhaps offsetting the fitness costs associated with increased sensitivity to GABA. In addition, the A2′S mutation conferred low levels of resistance to fipronil (IC50 45 nM compared to 20 nM in wild-type). However, in combination with R299Q, the sensitivity to fipronil was significantly decreased (IC50 96 nM), suggesting the double mutant also contributes to fipronil resistance. Therefore the A2′S and R299Q subunit mutations are likely to have both a compensatory and synergistic relationship; to offset fitness costs and increase fipronil resistance, respectively. 6

Table 1. Resistance Mutations Found in Association with the A2′ Mutation in the RDL Subunit of Various Insect Species. The Mutations Are Listed in Order of Position within the Protein Sequence, with the Insect Species Detected, Resistance Ratio if Calculated and Geographical Location from Which They Were Isolated, unless They Were Obtained via Laboratory Selection. RDL mutation associated with A2′

Insect species

Resistance ratio

Geographical location isolated

M2 domain A2′S and R299Q

N. lugens

Fipronil: 237

China, Vietnam, Thailand, plus via laboratory selection (67)

A2′S and R305Q or R305W

L. striatellus

Fipronil: 87

Laboratory selection (field collected strain from Japan) (66)

Loop between M2-M3 domains

7

A2′S and V327I

An. funestus

Not determined

Africa (Burkina Faso and Cameroon) (56)

M3 domain A2′G and T345M

An. gambiae

Not determined

Laboratory selection (field collected strain from Democratic Republic of Congo) (20)

A2′G and T350M

D. simulans (Eyguières 42)

Fipronil: 20,000

Laboratory selection (31)

Intracellular loop between M3-M4 domains A2′N and R340Q

S. furcifera

Not determined

Japan (65) Continued on next page.

Table 1. (Continued). Resistance Mutations Found in Association with the A2′ Mutation in the RDL Subunit of Various Insect Species. The Mutations Are Listed in Order of Position within the Protein Sequence, with the Insect Species Detected, Resistance Ratio if Calculated and Geographical Location from Which They Were Isolated, unless They Were Obtained via Laboratory Selection. RDL mutation associated with A2′

Insect species

Resistance ratio

A2′S and Q359E

N. lugens

Under ethiprole selection: Ethiprole: >14,000 Fipronil: >860

A2′S and M360I

D. melanogaster

Dieldrin: >4000

Geographical location isolated Laboratory selection (field collected strain (Nl55) from India) (30) USA (64)

8

Naturally-Occurring Variations: Alternative Splicing and RNA Editing Alternative splicing and RNA editing are post-transcriptional modifications that enhance the diversity of the transcriptome, thereby increasing the number of products from a single gene (70, 71). Ion channel proteins functioning in the nervous system undergo a notably high level of RNA editing, commonly in functionally significant regions, which may serve as a mechanism for facilitating rapid neuronal signaling (72). The importance of alternative splicing and RNA editing is highlighted by their dysregulation, which results in neurodegenerative phenotypes (73–75). These mechanisms may also be particularly influential in insect genomes, serving to expand the repertoire of what would otherwise be a small complement of proteins. For example, insects possess less than 30 cysLGIC subunit genes, as opposed to 102 in Caenorhabditis elegans and 45 in humans (27, 76, 77).

The Effects of Alternative Splicing on RDL Receptor Pharmacology Alternative splicing is the process in pre-messenger RNA (mRNA), whereby introns are spliced out and various exons are introduced, removed or substituted to form the final processed mRNA. In this way, a single gene codes for multiple proteins, as a result of the alternatively spliced exons forming multiple protein products (78, 79). D. melanogaster Rdl is composed of nine exons, two of which, exons 3 (variants a and b) and 6 (variants c and d) (Figure 1) are alternatively spliced to produce subunit isoforms ac, ad, bc and bd, all of which are transcribed in vivo (50, 51). Real time PCR and analyses of clones amplified from adult D. melanogaster and An. gambiae cDNA reveal that the Rdlbd variant is the most prevalent splice form in vivo (51, 80). However the preference for the splice variant transcript appears to be different at the embryonic stage, with Rdlbc being the most abundant (50), indicating varying requirement for the different isoforms at different stages of development. Alternative splicing of exons 3 and 6 is conserved in diverse insect species including the African malaria mosquito An. gambiae (80), the small brown planthopper L. striatellus (81), the silkworm B. mori (26), the honeybee A. mellifera (28), beetles T. castaneum and Oulema oryzae (82, 83) and the parasitoid wasp Nasonia vitripennis (27) also revealed these common splice regions. In contrast, the white-backed planthopper S. furcifera has an additional splice variant at exon 3 (65). The diamondback moth, P. xylostella, has two Rdl-encoding genes, both of which are alternatively spliced at only exon 3 (25). This suggests that even within insect orders, the level of alternative splicing can differ, creating species specific RDL subunit isoforms.

9

Figure 1. Schematic illustration showing how post-transcriptional modifications can diversify the insect RDL subunit, in this case that of D. melanogaster. Alternative splicing yields two variants each for exons 3 (variants a and b) and 6 (variants c and d) changing amino acid residues in the N-terminal extracellular domain where agonist binding occurs. Residues that are different in alternative exons are shown in bold. Differential splicing generates M3-M4 intracellular loops of different lengths as shown by sequences with the accession numbers NP_729462.2 and NP_523991.2 (sequences directly submitted to the NCBI database). RNA editing leads to the R122G substitution in the N-terminal extracellular domain, I283V in M1, whereas N294D and M360V occur in intracellular domains.

Exons 3 and 6 are located in the N-terminal extracellular domain, within proximity of agonist binding. In particular, exon 6 contains loops C and F, which contribute to the ligand-binding site (15). It was therefore not surprising to find that the splice isoforms in Drosophila differed in their sensitivity to GABA and various GABA analogues when expressed using the X. laevis expression system and two-electrode voltage clamp electrophysiology (51, 84–86). The sensitivity to GABA was in the order bc>ac>ad>bd, ranging in EC50 from 20 ± 0.2 µM (RDLbc) to 152 ± 10 µM (RDLbd) (51). More recently it has been shown that different subunit isoforms of RDL can also arise from variation in the large intracellular loop between M3 and M4. Here, instead of using alternative exons, the use of different splice acceptor sites generates intracellular loops of varying lengths (Figure 1). For example, the miridbug C. lividipennis contains two RDL subunit isoforms differing by a 31 amino acid insertion in the intracellular loop (23, 28). Two-electrode 10

voltage-clamp electrophysiology applied to C. lividipennis Rdl expressed in X. laevis oocytes showed that these two isoforms significantly differed in their sensitivity to fipronil, with the presence of the insertion significantly increasing the IC50 from 6.47 ± 1.12 µM to 16.83 ± 2.30 µM. This suggests that diversity in this large intracellular loop may enhance the tolerance to fipronil or other insecticides. Another insect species, A. mellifera, has three differentially spliced M3-M4 isoforms of RDL (22, 28). These variants, unlike those in C. lividipennis, did not differ in their sensitivity to fipronil (22). Interestingly, the site of insertion disrupts a putative protein kinase C phosphorylation consensus site, conserved in both A. mellifera, C. lividipennis and D. melanogaster RDL (22). Phosphorylation of the intracellular loop can influence events such as protein assembly, receptor desensitization and insecticide sensitivity (87–90). Therefore differential splicing has the potential to affect insecticide actions in a species specific manner. The differential effects of species specific subunit isoforms on insecticide sensitivity described above are highly pertinent, since species such as C. lividipennis and A. mellifera are both beneficial insects. This is highly relevant considering that the use of fipronil has been restricted by the European Union (91) for its suspected negative effects on bees. These findings hint at the complex level of species specific differential splicing that can occur in insects, which may impact on the response of receptors to insecticides. It remains to be seen whether differential splicing of the intracellular loop of RDL subunits are highly conserved in diverse insects, including pest species, and whether this presents another route to reducing sensitivity to insecticides. The Effects of RNA Editing on RDL Receptor Pharmacology RNA editing also diversifies the number of products produced from a single gene, but via single nucleotide substitutions, initiated by adenosine deaminases acting on RNA (ADAR) enzymes (92, 93). In the most prevalent form of RNA editing, known as A-to-I editing, ADAR enzymes deaminate adenosine to inosine, which is then translated as guanosine (94). The result is a sequence transcript dissimilar to that of the genomic DNA, potentially changing amino acid residues that may affect protein structure and function. For example, four RNA editing sites that alter amino acid residues were identified in clones amplified from D. melanogaster Rdl cDNA (51, 72) (Figure 1). In an analysis of >100 clones, the edit sites R122G, I283V, N294D and M360V were found in 16 different combinations, predominantly in the bd splice variant background (51). These amino acid substitutions are in structurally significant regions, making it likely that they will affect receptor function. For instance, R122G is located between ligand binding loops D and A, suggesting that it may influence agonist binding. Es-Salah et al. confirmed this finding by showing that when the arginine is replaced with glycine the GABA EC50 is significantly increased (52). Functional expression of the different editing isoforms also generated a range of sensitivities to GABA in D. melanogaster RDL (EC50s of 3-193 μM), showing that singly and in combination, the edit sites directly affect agonist potency (51). 11

Interestingly, the choice of splice variant arising from the alternative use of exons 3 and 6 influenced the potency of GABA in addition to RNA editing. This was shown by significantly different GABA EC50s for the ad and bd splice variants with the same edit combination, suggesting that editing and splicing may act in concert to further broaden the functional diversity of the RDL receptor (51). Resistance mutations in RDL subunits were also investigated in combination with RNA editing. The N-terminal R122G edit was investigated in comparison with an A2′G/T350M mutated RDL receptor isoform (52). The sensitivity to both GABA and fipronil was significantly reduced by the R122G addition, suggesting that RNA editing is able to affect insecticide sensitivity in RDL receptor isoforms bearing resistance mutations. The authors speculate that as fipronil acts preferentially on agonist-bound receptors and R122G reduces the potency of GABA, the reduction in the proportion of agonist bound receptors may affect fipronil binding (52). Lees et al. also investigated the effect of the RNA edit site I283V on fipronil sensitivity in A2′S mutant homomeric D. melanogaster RDL and found that this edit site did not affect GABA binding or fipronil sensitivity (36). This difference in effect may be due to the position of I283V in the M1 domain of the subunit, as it is not in close proximity to locations of agonist or antagonist binding. RNA editing can generate isoforms that are highly species specific. For example, one of the two Rdl genes (RDL 1) from B. mori generates transcripts with two potential RNA editing sites that alter two amino acid residues at the C-terminal end, a region completely different to those altered in D. melanogaster RDL (26). Recently, RNA editing sites were also found in the RDL subunit of various mosquito species, which are not completely conserved with D. melanogaster (80). In this study, a comprehensive sequence analysis was conducted on the RDL subunit from the mosquito disease vectors; An. gambiae, Culex pipiens and Ae. aegypti. Nine putative RNA editing sites were observed in Rdl cDNA sequences; five in Ae. aegypti, seven in Cx. pipiens and eight sites in An. gambiae. Two of these, I278V and N289D, were conserved in D. melanogaster (51). Therefore the remaining sites were not only mosquito specific, but also in some cases mosquito species specific. The functional effects of different RNA editing isoforms in RDL subunits from An. gambiae were investigated in the bd splice variant background (80). As with D. melanogaster, the edit combination generated a spectrum of potencies to GABA, with EC50s ranging from 5 ± 1 to 246 ± 41 μM in the 18 isoforms tested. Interestingly, the EC50 increased in line with an increase in the number of RNA edited sites. RNA editing did not however have an effect on the sensitivity of fipronil. This is perhaps unsurprising as none of the editing sites are in the pore-lining M2 region important for fipronil binding (15). The macrocyclic lactone ivermectin also acts on the GABA receptor, but the exact mechanisms of action are not clear. Ivermectin has been found to potentiate the GABA response of vertebrate GABA receptors (95) and the D. melanogaster RDL receptor (96) using patch clamp electrophysiology applied to cell lines and the FLIPR membrane potential assay. For M. domestica homomeric RDL receptors expressed in X. laevis oocytes, ivermectin was also found to potentiate currents induced by low concentrations of GABA (EC5) (97). However, 12

ivermectin has been shown to act as an antagonist on An. gambiae (80), D. melanogaster (36) and M. domestica (97) RDL currents, as homomeric receptors expressed in Xenopus oocytes, when induced by a GABA concentration higher than the EC50. RNA editing of An. gambiae RDL isoforms influences the potency of ivermectin, where it was shown to significantly reduce the IC50 of the unedited receptor from 457 ± 118 nM to 50 ± 24 nM in the completely edited isoform (80). Ivermectin was also found to directly activate the An. gambiae RDL receptor as well as potentiate the GABA-induced responses at concentrations below the EC50 (80). Differences between the edited and unedited isoforms were also observed under these experimental conditions. Concentrations of 0.01 μM and 0.03 μM ivermectin potentiated currents induced by GABA at the EC20 in the unedited receptor isoform, but not in the edited isoform (80). The triple actions (agonistic, potentiating and antagonistic) of ivermectin were also observed for the M. domestica RDL receptor (97) where the authors speculated that the number of orthosteric binding sites in the pentamer occupied by GABA, may determine whether ivermectin is a potentiator or antagonist. It remains to be seen whether RNA editing sites in An. gambiae RDL subunits contribute individually or in conjunction to elicit changes in the sensitivity to ivermectin. Of particular interest would be the N183G edit site of the RDL subunit from An. gambiae, which is within the cys-loop, and therefore could influence communication between the GABA binding regions and the transmembrane domain (98). This information may have relevance to controlling vector-borne diseases as ivermectin has been shown to reduce the longevity of An. gambiae mosquitoes (99). Intriguingly, not all insect species contain an RDL receptor that is RNA edited. For instance, RNA A-to-I editing sites have not been detected in RDL from the honey bee (A. mellifera), the red flour beetle (T. castaneum) and the parasitoid wasp (N. vitripennis) (27, 28, 82). RNA editing, therefore, can recode the highly conserved genomic sequence for Rdl in a species-specific manner. Why RDL subunits of some insect species undergo RNA editing and others do not remains a mystery. The finding that RNA editing generates numerous receptor isoforms of RDL in An. gambiae suggests that the mosquito GABA receptor requires a higher level of plasticity than that of the honey bee.

Conclusions and Future Prospects Identification of the dieldrin resistance mutation in the RDL receptor was pivotal to understanding the mechanism of target site resistance in insects. Dieldrin is no longer in use, however fipronil, which also shows varying levels of crossresistance to the RDL receptor, is used worldwide in various applications ranging from the control of agricultural pests to the treatment of various parasitic diseases (100). Therefore understanding variations in the structure and function of the RDL receptor is relevant to understanding mechanisms of resistance in the field, as well as a target for emerging novel classes of insecticides (39, 40). 13

Future investigations of the RDL receptor will likely continue to reveal further resistance associated mutations. The A2′S/G/N are the common variants of the classically conserved RDL mutation in M2, although a number of additional mutations have recently been identified in highly insecticide resistant insects (Table 1). In all instances, the second mutations were associated in parallel with the A2′ mutation. Identification of these mutations may indicate a gradual adaptive response to changing levels of insecticides in the environment. However, it may also be possible that these mutations have been present in populations for some time, but were not previously identified as the entire coding region of the Rdl gene is commonly not sequenced. To presume that the A2′ mutation is central to RDL receptor-based insecticide resistance may encourage diagnostic checks for resistance to concentrate solely on the M2 domain. This has the danger to miss other mutations that may be important (32). There is increasing evidence that post-transcriptional modifications can impact on RDL receptor pharmacology, providing another reason to sequence the whole coding region to identify subunit isoforms that may arise from alternative splicing and RNA editing. There is no doubt that in the future novel splicing and editing isoforms of the RDL receptor will continue to be identified as more cDNA clones are investigated. It would be additionally advantageous to further investigate the effects of splicing and editing in pest species in order to investigate their contribution as potential factors in altering the tolerance to insecticides. Future experiments with potentially new insecticides that act on the RDL receptor could include elucidating whether splicing and RNA editing affect the potency of these novel compounds. Can the findings of species-specific splicing and RNA editing of RDL subunits be exploited? Perhaps future insecticide discovery efforts may identify compounds preferentially acting on RDL receptor isoforms found only in pest species. This is considerably timely since the use of fipronil and neonicotinoids have been restricted amidst fears that they are having a detrimental effect on non-target organisms (100).

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Chapter 2

Insecticide Resistance in Rice Planthoppers Toshifumi Nakao* Agrochemical Research Center, Mitsui Chemicals Agro, Inc., Mobara, Chiba 297-0017, Japan *E-mail: [email protected].

Rice is a major crop in Asia. However, production of rice has been seriously damaged by three rice planthoppers: the brown planthopper (BPH), Nilaparvata lugens (Stål); the white backed planthopper (WBPH), Sogatella furcifera (Horváth); and the small brown planthopper (SBPH), Laodelphax striatellus (Fallén). To control rice planthoppers, various insecticides such as organochlorines, organophosphates, carbamates, pyrethroids, and buprofezin were used. However, rice planthoppers have developed resistance to these insecticides. In 1991, imidacloprid was introduced to control planthoppers. Imidacloprid suppressed the rice planthopper populations and was heavily used in Asia. However, populations of imidacloprid-resistant rice planthoppers have appeared. Since 2005, outbreaks of rice planthoppers have occurred in Asia. After introduction of imidacloprid, fipronil and ethiprole were commercialized to control rice planthoppers. But rice planthoppers have developed resistance to fipronil and ethiprole. In this chapter, insecticide resistance and its mechanisms in rice planthoppers are reviewed.

Introduction Rice is a major crop in Asia. However, production of rice has been seriously damaged by three rice planthoppers: the brown planthopper (BPH), Nilaparvata lugens (Stål); the white backed planthopper (WBPH), Sogatella furcifera (Horváth); and the small brown planthopper (SBPH), Laodelphax striatellus (Fallén).

© 2017 American Chemical Society

BPH attacks only rice plants, by sucking and transmitting viruses such as, rice ragged stunt virus and rice grassy stunt virus. WBPH causes sucking damage on rice plants. Recently, WBPH has been reported to cause southern rice blackstreaked dwarf virus disease. SBPH infests not only rice plants but also other crops, such as wheat, barely, and rye, and transmits viruses, such as rice stripe virus (1, 2). In Asia, BPH is found in Bangladesh, Brunei, Burma (Myanmar), China, regions of Hong Kong, India, Indonesia, Japan, Cambodia, Korea, Laos, Malaysia, Nepal, Pakistan, the Philippines, Singapore, Sri Lanka, regions of Taiwan, Thailand, and Vietnam (3). SBPH is found in Bangladesh, Cambodia, China, regions of Hong Kong, India, Indonesia, Japan, Korea, Laos, Malaysia, Myanmar, Nepal, Pakistan, the Philippines, Ryukyu Islands, Sri Lanka, regions of Taiwan, Thailand, and Vietnam (3). WBPH is observed in Bangladesh, China, India, Indonesia, Japan, Korea, Malaysia, the Philippines, Taiwan, Thailand, and Vietnam BPH and WBPH live in tropical regions of Southeast Asia. They emigrate from tropical regions of Southeast Asia into northern subtropical and temperate regions from May to July. But they do not overwinter in temperature regions, such as Japan and Korea. SBPH undergoes diapauses and overwinters in temperature regions. Furthermore, overseas migration of SBPH from China to western Japan was reported (4). To control planthoppers, various insecticides such as organochlorines, organophosphates, carbamates, pyrethroids, insect growth regulators, neonicotinoids, phenylpyrazoles, and pymetrozine, were used (5, 6). However, rice planthoppers have developed resistance to these insecticides. For example, the field collected populations of BPH in China from 2012 to 2014 had developed high levels of resistance to neonicotinoid insecticide imidacloprid (resistance ratio, RR = 233.3-2029) and buprofezin (RR = 147.0-1222). Furthermore, these populations of BPH showed low to moderate resistance to carbamte insecticide isoprocarb (RR = 17.1-70.2) and organophosphate insecticide chlorpyrifos (RR = 7.4-30.7) (7). There are sevelal insecticide resistant mechanisms, such as, target-site mutation, metabolic detoxification, penetration resistance, and excretion. In this chapter, insecticide resistance in rice planthoppers is reviewed.

Resistance to Organochlorines, Organophosphates, Carbamates, Pyrethroids, Buprofezin, and Pymetrozine Organochlorines, such as BHC (benzene hexachloride) and DDT (dichloro-diphenyl-trichloroethane), were the first insecticide used to control BPH (5). According to the Insecticide Resistance Action Committee (IRAC), BHC belongs to IRAC group 2A which acts as a GABA-gated chloride channel blocker. DDT belongs to IRAC group 3B which acts as modulators against sodium channels. Rice planthoppers developed resistance to BHC and DDT (8, 9). 24

Organophospahtes and carbamates belong to IRAC group 1 which acts as acethylcholinesterase (AChE) inhibitors. As organophospahtes and carbamates were widely used to control rice planthoppers, rice planthoppers developed resistance to these insecticides relatively fast (5, 8, 9). The main factor contributing to resistance of the BPH to organophospahtes and carbamates was suggested to be detoxification as a result of high esterase activity, which was caused by high expression of amplified carboxyesterase gene, Ni-EST1 (10–12). Comparison of amino acid sequences of the AChEs between the susceptible and resistant BPH strains revealed a point mutation, G185S. The G185S mutation was suggested to change the affinity of AChE for its substrates and inhibitors. Thus, G185S mutation was likely responsible for the insensitivity of the AChE to methamidophos in the resistant strain (13). Pyrethroids belong to IRAC group 3A and act as sodium channel modulators. Use of pyrethroids to control BPH was limited to only a few countries (5). Selection of a laboratory colony of the BPH with the pyrethroids permethrin and λ-cyhalothrin increased its resistance to both insecticides. Biochemical analysis and synergistic studies with metabolic inhibitors indicated that elevated glutathione S-transferases (GSTs) with a predominant peroxidase activity conferred resistance to both pyrethroids (14, 15). Etofenprox is a non-ester pyrethroid insecticide. Although the susceptibility of BPH to etofenprox remained a susceptible level of resistance in China (7), the successive selection by etofenprox for 16 generations in the laboratory resulted in a high level resistance and it was suggested that high expression of a P450 gene CYP6FU1 was associated with resistance (16). Buprofezin belongs to IRAC group 16 and acts as an inhibitor of chitin biosynthesis. Buprofezin was used from 1984 to control BPH, but the sensitivity of BPH to buprofezin began to decrease after 10 years (5). Most populations of WBPH in eastern China in 2010 and 2011 developed moderate resistance to buprofezin (up to 25-fold) (17). Pymetrozine is the first and only substance from the azomethyne pyridine (5). Pymetrozine belongs to IRAC group 9 and acts as a chordotonal organ TRPV channel modulator. Pymetrozine resistance of the nine field populations of the BPH collected from China in 2012 was at a moderate level. Resistance ratio of the nine populations ranged from 34.9 to 46.8-fold (18),

Resistance to Imidacloprid As mentioned above, rice planthoppers have developed resistance to organochlorines, organophosphates, carbamates, pyrethroids, and buprofezin. In 1991, imidacloprid was introduced to control planthoppers. Imidacloprid belongs to IRAC group 4A and acts as a nicotinic acetylcholine receptor (nAChR) agonist. Imidacloprid suppressed the rice planthopper populations and was heavily used in Asia. However, imidacloprid-resistant BPH was first observed in Thailand and appeared in Asian counties, such as, Vietnam, China, and Japan (19). Since 2005, outbreaks of BPH have occurred in East Asian countries (5). 25

The main resistant factor that confers imidacloprid resistance in BPH was suggested to be detoxification caused by increased P450 monooxygenase activity (20, 21). Imidacloprid-resistant BPH strain, which originally collected from a field population and continuously selected in laboratory with imidacloprid for more than 40 generations, had 180.8-fold resistance to imidacloprid, compared to a susceptible strain. Expression levels of CYP6AY1 mRNA was found to be highest in imidacloprid-resiatant BPH strain. By expressing CYP6AY1 in Escherichia coli cells, CYP6AY1 was found to metabolize imidacloprid efficiently. When CYP6AY1 mRNA levels in imidacloprid-resistant strain were reduced by RNA interference, imidacloprid susceptibility was recovered. In four field populations with different resistance levels, high levels of CYP6AY1 transcript were also found (22). In contrast, overexpression of CYP6ER1 was associated with field-evolved resistance to imidacloprid in BPH populations in five countries in South and East Asia (23). RNA interference of CYP6ER1 and transgenic expression of CYP6ER1 in Drosophila melanogaster both suggested that the expression of CYP6ER1 was sufficient to confer imidacloprid resistance (24, 25). It was suggested that four P450 genes (CYP6AY1, CYP6ER1, CYP6CS1 and CYP6CW1) conferred imidacloprid resistance to BPH and that CYP6ER1 was important at all stages of resistance development (25). Although target site mutation Y151S in two nAChR subunits from laboratory selected imidacloprid-resistant BPH was reported (26), this mutation has never been observed in any field collected populations (5). Continuous selection of a filed collected strain of BPH with imidacloprid in the laboratory resulted in a substantial increase in resistance and it was suggested that reduction in mRNA and protein expression of a nAChR α8 subunit was associated with resistance (27). SBPH also has developed resistance to imidacloprid in China, and immigration of SBPH into Japan from China caused imidacloprid resistance in Japan (4, 28). The main factor that confers imidacloprid resistance in SBPH was suggested to be detoxification caused by increased P450 monooxygenase activity of CYP353D1v2. Expression level of CYP353D1v2 of imidacloprid-resistant SBPH was significantly different to that of the susceptible strain. Strong correlation was found between CYP353D1v2 expression levels and imidacloprid treatments. Additionally, depression of the expression of CYP353D1v2 by RNA interference could significantly enhance the sensitivity of SBPH to imidacloprid (29).

Resistance to Fipronil Followed by introduction of imidacloprid, fipronil was commercialized to control rice planthoppers. Fipronil belongs to IRAC group 2B and acts as an RDL GABA receptor blocker (Figure 1). Fipronil was effective against rice palnthoppers and became one of the main alternatives after rice palnthoppers developed resistance to imidacloprid (5, 19). However, fipronil-resistant rice palnthoppers have been reported. 26

Figure 1. Structures of phenylpyrazoles, fipronil and ethiprole.

Figure 2. RDL GABA receptor subunit and mutations that confer fipronil resistance against planthoppers. SBPH has developed resistance to fipronil in Japan (19, 30). Sequence analysis of the Rdl genes from a fipronil-resistant SBPH population collected in Fukuoka Prefecture in Japan during 2009 identified an A2′N mutation (index number for M2 membrane spanning region) (Figure 2) in the heterozygous state (31). A membrane potential assay was carried out using Drosophila S2 cells expressing the wild-type and A2′N mutant SBPH Rdl genes, either individually or together. The EC50 value of GABA for the wild-type homomers was 0.72 μM. Expression of A2′N mutant receptor homomers decreased the sensitivity 27

to GABA. The EC50 value of GABA for the A2′N mutant receptor homomers was 11.0 μM (Figure 3). By contrast, the EC50 value for GABA in cells that expressed the wild-type and A2′N mutant Rdl genes was 1.4 μM (Figure 3), thereby indicating that co-expression of wild-type and A2′N mutant RDL GABA receptor subunits restored the sensitivity to GABA. The A2′N mutation abolished the inhibitory activity of fipronil in cells expressing the A2′N mutant Rdl gene with or without the wild-type Rdl gene (Figure 4) (31). A two-electrode voltage clamp study showed that the A2′N mutant from Musca domestica GABA receptors expressed in Xenopus oocytes decreased the sensitivity to fipronil profoundly (32). A fipronil-resistant SBPH population selected in a laboratory showed 112.1-fold resistance to fipronil and 24.5-fold resistance to ethiprole. A2′N mutation was suggested to play an important role in conferring both fipronil and ethiprole resistance (33, 34). To predict the appearance of fipronil-resistant SBPH, a rapid polymerase chain reaction-restriction fragment length polymorphism (PCR-RFLP) assay has been developed (35). One hudred and fourteen individuals of SBPH were collected from 4 regions in Gunma Prefecture in Japan in 2014 and more than 80 % of SBPH individuals were diagnosed as insects carrying A2′N mutation in the Rdl gene using the PCR-RFLP assay (36). Wei et al. (2016) has developed the ASPCR assay to detect the A2′N mutation in the SBPH Rdl gene, and the mutation frequencies of 19.2% and 3.6% had appeared in Lujiang and Gaochun populations in China in 2016, respectively (34).

Figure 3. Concentration–response curves for GABA in the SBPH RDL GABA receptors. Data are expressed as percentages of the maximal response to GABA for homo-oligomeric wild-type SBPH RDL GABA receptor homomers (open circle), homo-oligomeric A2′N mutant SBPH RDL GABA receptors (open square), and RDL GABA receptors on cells co-transfected with wild-type and A2′N mutant SBPH Rdl genes (open triangle). Vertical bars represent SEM for three independent experiments done in duplicate. (Reproduced with permission from ref. (31). Copyright 2011 Oxford University Press). 28

Figure 4. Concentration–response curves for fipronil in the SBPH RDL GABA receptors. Data are expressed as percentage inhibition of the response to EC80 concentrations of GABA in the absence of fipronil with homo-oligomeric wild-type SBPH RDL GABA receptors (open circle), homo-oligomeric A2′N mutant SBPH RDL GABA receptors (open square), and RDL GABA receptors on cells co-transfected with wild-type and A2′N mutant SBPH Rdl genes (open triangle). Vertical bars represent SEM for three independent experiments done in duplicate. (Reproduced with permission from ref. (31). Copyright 2011 Oxford University Press). WBPH has also developed fipronil resistance (19, 37). Biochemical studies indicated that the esterase and P450 monooxygenase might be related to fipronil resistance, but the existence of other more important factors have been suggested (37). From a fipronil-resistant WBPH population collected in Japan in 2007, the A2′N mutation was found in the heterozygous state (Figure 2) (38). In addition, a novel R340Q mutation was associated with the A2′N mutation in the cytoplasmic loop M3–M4 (Figure 2) (39). A membrane potential assay showed that the EC50 value for GABA with the wild-type receptor homomer was 0.25 μM. The EC50 values for GABA with A2′N and A2′N · R340Q mutant receptor homomers were 5.63 and 6.03 μM, respectively (Figures 5A and 6A, respectively) (39). As is the case for SBPH RDL GABA receptors, the decreased sensitivity of the A2′N and A2′N · R340Q mutant WBPH RDL GABA receptors was recovered by coexpression of the wild-type receptor (Figures 5A and 6A, respectively) (39). The IC50 value for fipronil with the wild-type receptor homomer was 79 nM (Figure 5B) (39). The RDL GABA receptors were inhibited by up to 40% with 3 μM fipronil in cells that expressed the wild-type and A2′N mutant genes (Figure 5B). By contrast, A2′N · R340Q double-mutant gene abolished the inhibitory activity of fipronil in cells that expressed the wild-type and A2′N · R340Q double-mutant genes (Figure 6B) (39). These results suggest that the A2′N · R340Q double mutation confers a higher level of resistance to fipronil than the A2′N mutation in the heterozygous state (Figures 5B and 6B, respectively) (39). 29

A rapid PCR-RFLP assay was also developed to detect the fipronil-resistant WBPH carrying the A2′N mutation in the Rdl gene (35). The PCR-RFLP assays for SBPH and WBPH are useful for monitoring fipronil resistance in planthoppers that carry the A2′N mutation in the Rdl gene, and they may facilitate the management of fipronil-resistant planthoppers.

Figure 5. Influence of the A2′N mutation on the GABA and fipronil sensitivities in WBPH RDL GABA receptors. (A) Concentration–response curves for GABA. Data are expressed as percentages of the maximal response to GABA with wild-type WBPH RDL GABA receptor homomers (open circle), A2′N mutant WBPH RDL GABA receptor homomers (closed square), and RDL GABA receptors on cells co-transfected with wild-type and A2′N mutant WBPHRdl genes (open square). Vertical bars represent SEM for three independent experiments done in duplicate.(B) Concentration–response curves for the inhibitory effects of fipronil on wild-type WBPH RDL GABA receptor homomers (open circle), A2′N mutant WBPH RDL GABA receptor homomers (closed square), and RDL GABA receptors in cells co-transfected with wild-type and A2′N mutant WBPH Rdl genes (open square). The inhibitory effects of fipronil on the response to EC80 concentrations of GABA are shown as the percentage difference relative to control cells in the absence of fipronil. Vertical bars represent SEM for three independent experiments done in duplicate. (Reproduced with permission from ref. (39). Copyright 2013 the Pesticide Science Society of Japan). 30

Figure 6. Influence of the A2′N• R340Q double mutation on the GABA and fipronil sensitivities of the WBPH Rdl GABA receptors. (A) Concentration–response curves for GABA. Data are expressed as percentages of the maximal response to GABA with A2′N•R340Q mutant WBPH RDL GABA receptor homomers (closed square) and RDL GABA receptors on cells co-transfected with wild-type and A2′N•R340Q mutant WBPH Rdl genes (open square). For comparison, the concentration–response curve for GABA with wild-type WBPH RDL GABA receptor homomers is also represented by a line. Vertical bars represent SEM for three independent experiments done in duplicate. (B) Concentration–response curves for the inhibitory effects of fipronil on A2′N•R340Q mutant WBPH RDL GABA receptor homomers (closed square) and RDL GABA receptors on cells co-transfected with wild-type and A2′N•R340Q mutant WBPH Rdl genes (open square). For comparison, a concentration–response curve for fipronil with wild-type WBPH RDL GABA receptor homomers is also represented by a line. The inhibitory effects of fipronil on the response to EC80 concentrations of GABA are shown as the percentage different relative to control cells in the absence of fipronil. Vertical bars represent SEM for three independent experiments done in duplicate. (Reproduced with permission from ref. (39). Copyright 2013 the Pesticide Science Society of Japan)

In contrast to SBPH and WBPH, fipronil-resistant BPH has not been observed until recently (30). BPH populations collected from six field populations in China in 2009 showed fipronil resistance with a 23.8-to 43.3-fold resistance ratio (40). Fipronil-resistant BPH was selected in laboratory. As the generation increased, 31

frequency of A2′S mutation (Figure 1) in the Rdl gene increased and R0′Q · A2′S double mutation (Figure 1) appeared. R0′Q · A2′S double mutation caused a much higher level of fipronil resistance (41). In field populations from China, Vietnam and Thailand, A2′S mutation was detected. Furthermore R0′Q · A2′S double mutation was observed in field populations from China and Vietnam, although the frequency of the double mutation was low. These results showed that fipronil resistance and related target mutations were widely distributed in field populations, but the low frequency indicated that the target mutations were not the dominant mechanism for fipronil resistance in the field populations (41).

Resistance to Ethiprole Ethiprole, the structure of which is similar to fipronil, also belongs to IRAC group 2B and acts as an RDL GABA receptor blocker (Figure 1). Unfortunately, resistance to ethiprole in BPH has been reported in China, Thailand, Vietnam, and India (23, 40, 42). An ethiprole-resistant population collected from Thailand developed 308.5-fold resistance to ethiprole and further selection with ethiprole for nine generations led to 453.1-fold resistance. Synergism and biochemical studies have suggested that the esterase and P450 monooxygenase activities caused the ethiprole resistance, although involvement of mutations in GABA receptor was also speculated (42). Garrood et al. (43) showed a casual cause of A2′S mutation on the GABA receptor in ethiprole resistance. Two ethiprole-resistant populations collected from Thailand and India showed 406- and 331-fold resistance, respectively. In contrast, fipronil resistance ratios of BPH populations collected from Thailand and India were 32-fold and 3-fold, respectively (43). After selection of these strains with ethiprole, ethiprole resistance ratios of these strains increased to >14,000-fold and fipronil resistance ratios of these strains reached 860-fold. The ethiprole-selected population from India showed 100% homozygous for A2′S mutation. Thus, major factor of mechanism of ethiprole resistance in BPH seems to A2′S mutation. SBPH also has developed ethiprole resistance (44). The resistance ratio of an ethiprole-resistant population of SBPH collected in China in 2013 was 107-fold and increased to 180-fold after selection with ethiprole. P450 monooxygenase genes, CYP4dE1 and CYP6CW3v2 were overexpressed in the resistant strain. When mRNA levels of CYP4dE1 and CYP6CW3v2 in ethiprole-resistant strain were reduced by RNA interference, ethiprole susceptibility was recovered, suggesting that CYP4dE1 and CYP6CW3v2 play an important role in ethiprole resistance in SBPH (44). Although approximately 25% of ethiprole-resistant SBPH carried the A2′N mutation, mutation might not be the major mechanism of observed ethiprole resistance. It is possible that frequency of A2′N mutation increase if selection continues (44).

32

Effects of A2′ Mutations in Drosophila melanogaster RDL GABA Receptors on Inhibitory Activities of Phenylpyrazoles We examined the effects of A2′ mutations in Drosophila melanogaster RDL GABA receptors on inhibitory activities of fipronil and ethiprole. Three different concentrations of GABA, EC50, EC80, and EC95 of GABA, were used, because a concentration-dependent effect of GABA on the insensitivity to fipronil in the A2′S mutant Oulema oryzae RDL GABA receptor was suggested (45). The membrane potential assay showed that there was no difference in fipronil sensitivity between the wild-type and A2′S mutant receptor homomers when EC50 and EC80 of GABA were applied (Figures 7A and 7B, Table 1). However, an approximately 14-fold reduction in the potency of fipronil was observed with the A2′S mutant receptor homomers when EC95 GABA was applied. By contrast, the fipronil sensitivity of the wild-type receptor homomers was not affected by the application of EC95 of GABA (Figures 7A and 7B, Table 1), suggesting that inhibitory activity of fipronil was affected by GABA concentration in A2′S mutant receptor.

Table 1. Comparison of Inhibitory Activities of Fipronil and Ethiprolea Fipronil

Wildtype

A2′S

A2′N

A2′G

Ethiprole

GABA concentration

IC50 (nM)

pIC50

IC50 (nM)

pIC50

EC50= 1.0 μM

69.8

7.156±0.054

55.7

7.254±0.041

EC80= 2.0 μM

33.7

7.472±0.061

53.9

7.268±0.033

EC95= 5.0μM

34.4

7.464±0.058

327.4

6.485±0.048

EC50= 1.4 μM

60.4

7.219±0.052

465.5

6.332

EC80= 3.0μM

42.3

7.373±0.036

3565.0

5.448

EC95= 8.0 μM

822.0

6.085±0.056

>10000

10000

10000

10000

10000

10000

10000

50% of currently marketed drugs (16). Cellular location, physiological function, and abundance in the cellular membrane, are among additional factors that make human GPCRs “druggable” targets, and these receptors are under wide investigation for development of novel therapeutics. Arthropod GPCRs regulate many biological processes, including reproduction, osmoregulation, growth, development and behavior (34). These receptors are presumed to exhibit a similar level of complexity to vertebrate GPCRs in terms of ligand binding and signaling modes, yet relatively less is known regarding their pharmacology. Arthropod GPCRs have been proposed as candidates for the development of next generation pesticides since the modification of receptor function by blocking or over stimulating its actions may either result in the death of a pest or disrupt its normal fitness and/or reproductive capacity, leading to a reduction in population number and disease transmission. Theoretically, arthropod GPCRs could be targeted by synthetic small molecule and natural product formulations that act either as agonists or antagonists. Alternatives include RNA interference (RNAi)- or Crispr/Cas9-based technologies that reduce or eliminate receptors that are essential to life. Target-driven insecticide discovery is enjoying renewed interest. The availability of sequenced and annotated invertebrate genomes has facilitated the identification of thousands of gene models, and in conjunction with techniques such as proteomics, functional genomics, and knockdown technologies, has provided information on essential physiological processes that could be exploited for pest control (34). Genome assemblies for multiple species of vectors, 60

including mosquito vectors of malaria, arboviruses and filariasis, the tsetse fly vector of African trypanosomiasis, sand fly vectors of leishmaniasis, kissing bug vector of Chagas, and an ixodid (hard) tick vector of Lyme disease, anaplasmosis and babesiosis are available via the National Institutes of Health (NIH) funded Bioinformatics Resource Center, VectorBase (3, 35). In 2011, the i5k initiative was launched with the goal to sequence the genomes of 5,000 insects and other arthropods considered of global importance for agriculture, food security, medicine and energy productions (36). These and other “omics” initiatives are expected to provide an unparalleled resource for target identification and comparative studies of orthologous targets, including GPCRs, across a wide range of phyletic groups. Numerous studies have proposed GPCRs as targets for insecticide development (19, 20, 23, 34, 37–40). Investigations have emphasized Class A receptors that bind small molecule biogenic amines, neuropeptides and peptide hormones. Formamidines are the only insecticide class recognized by IRAC that operate via disruption of GPCR function (9). The formamidine, amitraz and its metabolite dimethylphenylmethyformamidine (DPMF) are reported to agonize the presumably arthropod specific α- and β-adrenergic-like octopamine receptors (40, 41). The search for novel control compounds that target octopamine and dopamine receptors has received significant interest, and the former target class is the subject of a recent review (42). Genome-wide RNA interference (RNAi)-based transcript knockdown studies provided target validation of dopamine and serotonin receptors in T. castaneum. Short interfering-RNA (siRNA) screens identified six GPCRs required for either larval development or ecdysis (43). Work by Regna et al. (44) using transgenic D. melanogaster provided validation of the D1-like dopamine receptor, DOP2. RNA interference (RNAi) knockdown of DOP2 transcripts resulted in significant fly lethality and results suggested a link to immune function and regulation of ecdysis.

Advances in Understanding of the Pharmacology of Arthropod GPCRs Efforts to characterize the pharmacology of arthropod receptors have largely focused on members of the biogenic amine-binding Class A receptors (17), with an emphasis on dopamine (DAR), octopamine/tyramine (OAR/TAR) and serotonin (5HT) receptors (Table 1). Biogenic amine receptors have proven amenable to cloning and expression in heterologous cell-based systems and functional assays are available to investigate receptor response to a variety of ligands. Notable studies include work on receptors from species of locust, silkworm, honey bee, fruit fly, cockroach, crickets, mosquitoes and ticks (38, 41, 45–52). Biogenic amines are derivatives of aromatic amino acids, and regulate a variety of behavioral and physiological processes, including locomotion, aggression, circadian rhythm, cardio-vascular control, learning and memory in invertebrates (53, 54). In contrast, the systems biology of biogenic-amine signaling in arthropods is a field in its infancy. Biogenic amines are also important 61

chemical messengers during embryonic and larval development, and synaptic organization of the brain. Dopamine and serotonin play fundamental roles in modulation of salivary gland function, reproduction, developmental (55–57), learning and memory (58), and diuresis (59). Octopamine and tyramine, the decarboxylation products of tyrosine, are functional counterparts of the vertebrate adrenergic transmitters epinephrine and norepinephrine (54). Octopamine plays key roles in modulating muscle activity in locusts, learning and memory in honeybees and fruit flies and stress response in crickets (60). The invertebrate octopaminergic system has received considerable research attention; detection of trace amounts of octopamine in mammals (58) has lead to speculation that the pathway is unique to arthropods. Notable studies include work on membrane preparations, neurons and cloned receptors in a number of invertebrate species (41, 44, 46, 49, 61–63). Octopamine functions in insects as a neurotransmitter, neurohormone and neuromodulator (20) and interacts with the invertebrate “α-adrenergic-like” (OctaR), “β-adrenergic-like” (OctbR), and “octopamine/tyramine” receptors (Oct/TyrR) which were identified based on their similarities in structure and in signaling properties to vertebrate adrenergic receptors (64). Pharmacological characterization of OARs and TARs is performed using agonists and antagonists identified from studies of the mammalian adrenergic system. Some progress has been made towards the identification of novel chemical entities active at vector OAR targets. Six candidate OAR/TAR genes were identified in the genome of the malaria mosquito, Anopheles gambiae and two OARs were cloned and functionally characterized by Kastner et al (49). Subsequent virtual screening using a homology model for one OAR enabled the identification of several agonist and antagonists, some of which exhibited toxicity to mosquito larvae. While octopamine receptor(s) are considered the primary target of formamidine metabolites, questions remain as to the role of OAR and TAR receptors in determining insecticidal effect. The interplay between OARs and TARs in response to formamidines has been investigated in the Bombyx mori (silkworm) system (41, 45, 46, 65–68). Functional assays involving the B. mori α-adrenergic-like OAR suggest that elevation of intracellular cAMP rather than Ca2+ mobilization might account for the insecticidal effect of formamidine insecticides (68). The B. mori β-adrenergic-like OAR showed a concentration-dependent increase in cellular cAMP in response to octopamine and a biphasic response to dimethylchlordimeform (46) in functional assays, while the tyramine receptor, TYR expressed in HEK-293 revealed an effect of BTS-27271 and demethylchlordimeform on forskolin-stimulated cAMP when the receptor was activated by tyramine (53). Similarly, pharmacological characterization of a putative OAR from the southern cattle tick, Rhipicephalus (Boophilus) microplus suggested that the receptor was likely a type-1 tyramine receptor (TAR-1) (69). Functional assays using heterologous expression of the receptor in Chinese hamster ovary cells (CHO-K1) showed strong fold-potency of tyramine versus octopamine at the receptor, antagonism by the α2-adrenergic antagonists, yohimbine and cyproheptadine, and agonistic effect of the amitraz metabolite BTS-27271 in the presence of tyramine. 62

Table 1. Summary of Invertebrate Class A (rhodopsin-like) G Protein-Coupled Receptors Showing Numbers of Predicted Receptors Identified in Each Family and Sub-family Sub-Family

Anopheles gambiae23

Aedes aegypti24

Culex quinque fasciatus25

Drosophila melanogaster22

Ixodes scap ularis26

Biogenic amine

Dopamine

4

6

3

2

6

Serotonin

6

11

2

6

4

Muscarinic acetylcholine

2

3

1

2

2

Histamine

1

1

-

-

-

Melatonin

1

1

Octopamine/ Tyramine

4

6

4

2

4

Orphan

-

-

5

9

-

Total

18

28

15

21

16

12

11

11

7

7

1

2

1

1

3

3

-

4

-

63

Receptor Family

Opsins Purine Glycoprotein hormone

Adenosine

3

-

Continued on next page.

Table 1. (Continued). Summary of Invertebrate Class A (rhodopsin-like) G Protein-Coupled Receptors Showing Numbers of Predicted Receptors Identified in Each Family and Sub-family

64

Receptor Family

Sub-Family

Anopheles gambiae23

Aedes aegypti24

Culex quinque fasciatus25

Drosophila melanogaster22

Ixodes scap ularis26

Peptide

Allatostatin/ ACP/Allotropin/ Galanin

3

-

3

2

14

Leukokinin/ Neurokinin/ Tachykinin

5

9

5

5

22

GH secretagoge /Neurotensin/ TRHR/LH/ FSH/TRH

2

2

-

7

-

Gastrin/ Bombesin/CCK

4

8

1

4

-

GnRH

3

3

1

2

-

Neuropeptide Y/F

4

11

2

5

2

Bursicon/Opioid

1

-

1

1

Somatostatin

1

1

2

2

-

Vasopressin

2

2

1

1

-

Unclassified

-

3

10

4

22

Total

25

39

25

33

61

Receptor Family

Anopheles gambiae23

Aedes aegypti24

Culex quinque fasciatus25

Drosophila melanogaster22

Ixodes scap ularis26

Orphan Class A

21

20

-

14

43

TOTAL

80

103

52

80

130

Sub-Family

ACP, adipokinetic/corazonin-related peptide; CCK, cholecystokinin; FSH, follicle stimulating hormone; GH, growth hormone; GnRH, gonadotropin-releasing hormone; 5HT, 5-hydroxytryptamine; LH, luteinizing hormone; TRHR, thyroid releasing hormone receptor; TSH, thyroid stimulating hormone; MAchR, muscarinic Acetylcholine.

65

Vertebrate DARs are classified as either D1-like (Gαs-coupled) or D2-like (Gαi-coupled) according to G protein coupling mechanisms (Figure 1). The neurotransmitter dopamine has important functions both in the central and peripheral nervous systems of vertebrates (70). Research has emphasized understanding and treatment of multiple neurological afflictions involving dopaminergic processes, including drug addiction, Parkinson’s disease and schizophrenia (71, 72). DAR-mediated regulation of intracellular cAMP is important for controlling signal transduction cascades triggered by phosphorylation activities of cAMP-dependent protein kinase (37). There are numerous reports involving cloning-based pharmacological characterization of invertebrate GPCRs, including D1- like and D2-like DARs from insects (38, 48, 50, 66, 73–76). Interestingly, D1-like DARs, and elevated cAMP levels have been implicated in tick salivation, a process linked to pathogen transmission during blood feeding (77, 78). Mustard et al (74) showed that inhibition of dopamine signaling decreased activity level in honeybees. This study provides a foundation for future work examining the importance of dopamine signaling in regulating distinct behaviors and elucidating the roles of specific dopamine receptors. Progress has been made towards an understanding of the pharmacology of mosquito and tick dopamine-binding receptors (DARs). Molecular and pharmacological studies identified two D1-like DARs in each of the mosquitoes Aedes aegypti, An. gambiae and Culex quinquefasciatus and the tick Ixodes scapularis (38, 39, 48, 50, 76). The DARs were identified based on genome assemblies available at NCBI and curated by VectorBase (35). Pharmacological studies revealed Gαs-coupling of receptors in vitro, and receptor agonism by dopamine and inhibition via a variety of small molecule antagonists (Figure 2). Importantly, lead elaboration combined with in vitro and in vivo SAR identified potent antagonists of the Ae. aegypti DOP2 DAR (79), enabling development of a preliminary pharmacophore. Some of these molecules exhibited two-fold selectivity for mosquito DARs versus the human ortholog, hD1 (Table 2). Additionally, Hill et al (76) showed pleiotropic coupling of the mosquito DARs expressed in a heterologous system to both Gαs and Gαq, highlighting possible plasticity in signaling modes. The conserved sequence and pharmacology of DARs between culicine (Aedes and Culex species) and anopheline (Anopheles species) mosquitoes suggests potential to develop products effective against multiple vectors and residual transmission by multiple anopheline vectors (76). Parallel characterization of dopamine receptors is an important step toward understanding the biological roles of dopaminergic processes in mosquitoes and ticks and generation of a pipeline for high throughput chemical screening of stably expressed vector GPCRs.

66

Figure 2. Pharmacological characterization of the Aedes aegypti AaDOP1 and AaDOP2 receptors. The mosquito receptors were stably expressed in HEK 293-CRELuc cells for dose-response assays. A, C: AaDOP1, B, D: AaDOP2. Representative curves for A, B: biogenic amines; C, D: synthetic dopamine receptor agonists; E: Inhibitory effect of 10 µM SCH23390 in the presence of 1 µM dopamine (n=4) shown for both mosquito dopamine receptors. ** p OA >> 5-HT.

Figure 4. Effects of biogenic amines on Ca2+-dependent luminescence in HEK-mock and HEK-BmDopR2 cells. HEK-BmDopR2 cells were transiently transfected with pcDNA-cytAEQ. Four typical biogenic amines were added at 10–3 M to the cells. HEK-mock cells were used as a negative control. Data represent the mean ± SE of three independent experiments, each done in duplicate. OA, octopamine; TA, tyramine; DA, dopamine; 5-HT, serotonin. (see color insert)

Effects of DA or OA Receptor Synthetic Agonists on Ca2+-Dependent Luminescence in HEK-BmDopR2 Cells To confirm whether the agonist profile of BmDopR2 can be discriminated from that of the phylogenetically related receptor BmOAR1, the typical nonselective vertebrate DA receptor agonist 6,7-ADTN, reported as the most effective BmDopR2 agonist in our previous study using cAMP assays (35), and the representative OA receptor agonists demethylchlordimeform (DMCDM) 114

(46) and NC-5 (47) were examined at 10–3 M, in parallel with DA, for their Ca2+-dependent luminescence response as agonist activity for BmDopR2 (Figure 5, black bars). In HEK-BmDopR2 transfected with pcDNA3-mtAEQ, 6,7-ADTN induced an approximately 60-fold increase in luminescence output relative to the control. The agonist activity at 10–3 M was comparable to that of DA. In contrast, DMCDM and NC-5 did not show remarkable agonist activity, although NC-5 induced a chlorpromazine > SCH-23390 > butacramol > spiperone (35). Thus, Flu is a potent BmDopR2 antagonist, effectively inhibiting both the receptor-mediated cAMP and the Ca2+ signals. However, whether Flu is also an effective antagonist for BmOAR1 as with the AmDOP2 and AmOA1 receptors remains to be examined. We are currently investigating the common pharmacological ground between BmDopR2 and BmOAR1 using a wide range of antagonists, including Flu. DMCDM is a well-known pesticide acting at OA receptors as an agonist (31, 46, 56). Accordingly, the compound has been reported to show agonist action for BmOAR1 (38, 57) and another B. mori OA receptor subtype, BmOAR2 (58). Here, we examined the agonist activity of DMCDM for BmDopR2; it had almost no agonist effect, but instead showed low (at least one orders of magnitude lower potency than that of Flu), dose-dependent antagonist activity. In contrast, the analogous agonist NC-5 did not show remarkable agonist and antagonist activities for BmDopR2 in this study, although at 10–3 M, it induced a marginal agonist effect. As for BmOAR1, however, this ligand is an agonist more potent than OA and equal to DMCDM at producing intracellular cAMP, and shows a sigmoidal intracellular Ca2+ response, with the rank order of potency of NC-5 > OA > DMCDM (57). Although paralogous to BmOAR1, BmDopR2 would have a unique pharmacology different from that of BmOAR1. 118

In this study, the luminescence produced by two types of apo-aequorin, cytAEQ and mtAEQ, was compared. As expected, HEK-BmDopR2 cells transfected with pcDNA3-mtAEQ displayed a higher DA-induced luminescence response than cells transfected with pcDNA3-cytAEQ, whereas the potency of DA was substantially equal. This result indicates that the mtAEQ-based assay would be more suitable to detect a weak Ca2+ signal mediated by GPCRs expressed at a low level on the cell membrane. Using the present aequorin assay system or the 96-well plate-based aequorin assay reported by Lu et al. (59) for functional analysis of the mosquito and tick neuropeptide kinin receptors, we plan to screen agonists and antagonists acting at BmDopR2 that should facilitate the development of lepidopteran-specific insecticides or insectistatics, because DopR2 receptors have been recently reported as potential targets for pest insect control (27, 29, 30, 60). In addition to BmDopRs, a B. mori peptide hormone GPCR, BmETHR, was expressed and functionally examined in HEK-293 cells using the aequorin assay. We could successfully detect BmETH-dependent Ca2+ signaling mediated by BmETHR. Considering this result together with other successful cases (BmOAR1 (42) and elevenin receptor NlA42 (43)), this aequorin assay may be useful as an efficient screening system for new control chemicals targeting biogenic amine and neuropeptide GPCRs in a wide array of pest insects.

Materials and Methods Chemicals DA·HCl was purchased from Nacalai Tesque (Kyoto, Japan). TA·HCl was from Wako Pure Chemical Industries (Osaka, Japan). OA · HCl was obtained from Aldrich (Milwaukee, WI, USA). 5-HT·HCl, 6,7-ADTN ((±)-2-amino-6,7-dihydroxy-1,2,3,4-tetrahydronaphthalene) · HBr, and cis-(Z)-flupenthixol·HCl were from Sigma (St. Louis, MO, USA). DMCDM and NC-5 (2-(2,6-diethylphenylimino)imidazolidine) were gifts from Dr. Y. Ozoe (Shimane University, Japan). Coelenterazine (native) was purchased from LUX Biotechnology (Edinburgh, UK). Other general reagents were purchased from Nacalai Tesque or Wako. BmETH was donated by Dr. H. Satake (Suntory Foundation for Life Sciences, Japan). Expression Vectors of BmDopRs and BmETHR As described in our previous study (3, 35), ORFs with stop codons of BmDopR1 and BmDopR2 cDNAs (accession nos. AB362162 and AB162716) were ligated into the BamHI and the KpnI/XbaI sites of the expression vector pcDNA3 (Invitrogen, Carlsbad, CA, USA), respectively, to produce pcDNA3-BmDopR1 and -BmDopR2. The ORF fragment of BmDopR3 cDNA (accession no. LC228577) was inserted into the KpnI/XbaI sites of pcDNA3 to produce pcDNA3-BmDopR3. 119

To construct the expression vector for BmETHR, pcDNA3-BmETHR, the ORF fragment of BmETHR cDNA, which was originally identified by comprehensive cloning of B. mori neuropeptide GPCR genes (45), was ligated into the KpnI/XbaI sites of pcDNA3. Cell Culture and Stable Transfection HEK-293 cells were grown in Dulbecco’s modified Eagle’s medium (DMEM; Invitrogen) containing 100 units/ml penicillin and 100 μg/ml streptomycin (Invitrogen) supplemented with 10% fetal bovine serum (FBS; Invitrogen) at 37°C and with 5% CO2. In our previous study, HEK-BmDopR1 and -BmDopR2 cells were constructed as polyclonal cell populations by stable transfection with pcDNA3-BmDopR1 and -BmDopR2, respectively (3, 35). Similarly, HEK-BmDopR3 and -BmETHR polyclonal cells were constructed. HEK-mock cells, polyclonal cells followed by stable transfection with empty pcDNA3 (3, 35), were used as negative control cells in aequorin Ca2+ assays. These five polyclonal cell lines were stored in medium-DMSO (Invitrogen) at –80°C until use. RT-PCR To Confirm Stable Expression of BmDopR1–3 in HEK-293 Cells Total RNA was extracted from HEK-mock, and HEK-BmDopR1–3 cells and used for single-stranded cDNA synthesis as described previously (61). PCR to confirm stable expression of each silkworm DA receptor in HEK-293 cells was carried out using the respective single-stranded cDNAs as templates and the following primer pair designed to target sites adjacent to the T7 and Sp6 promoter regions in pcDNA3 (forward, T7-F: 5´-TAATACGACTCACTATAGGG-3´; reverse, Sp6-R: 5´-CTCTAGCATTTAGGTGACAC-3´). The thermal cycles were as follows: denaturation at 94°C for 5 min, 23 cycles of 0.5 min at 94°C, 1 min at 51°C and 2 min at 72°C, and a final extension at 72°C for 5 min. In the same PCR batch, a region of β-actin cDNA as a reference gene was amplified using the cDNA templates used above and 5´-CTGAAGTACCCCATCGA-3´ as a forward primer and 5´-CATGATCTGGGTCATCTT-3´ as a reverse primer. PCR products were electrophoresed on a 1% agarose gel in 1× TAE. Construction of Aequorin Vectors Apo-aequorin cDNA cloned in pIZT/V5-His (Invitrogen) was donated by Drs. T. Tanimura and H. Ishimoto (Kyushu University, Japan). The ORF region with KpnI and XbaI sites attached to the 5´- and 3´-terminals, respectively, was amplified by standard PCR using the DNA polymerase KOD-Plus- (Toyobo Co., Ltd., Osaka, Japan), the aequorin vector, pIZT/V5-His-AEQ, as a template, and the following primer pair (forward: 5´-ATATGGTACCATGACAAGCAAACAA-3´; reverse: 5´-ATATTCTAGATTAGGGGACAGCTC-3´). The amplified product was treated with the restriction enzymes and ligated into the corresponding sites of pcDNA3 to produce pcDNA3-cytAEQ. To produce pcDNA3-mtAEQ, a mitochondrial targeting signal sequence was derived from the cDNA sequence encoding the N-terminal 33 amino acid residues of subunit VIII of human 120

cytochrome c oxidase. The segment was amplified by standard PCR using KODPlus-, with single-strand cDNA synthesized from HEK-mock cells as a template and 5´-ATATAAGCTTATCATGTCCGTCCTG-3´ with a HindIII site as a forward primer and a Kozak sequence and 5´-ATATAAGCTTCCCCTCCGGCGGCAA-3´ with a HindIII site as a reverse primer. The obtained fragment was treated with the restriction enzyme and inserted into the two HindIII sites of pcDNA3-cytAEQ; one remained in the multi-cloning site and the other was located at nucleotides 25–30 of the apo-aequorin ORF region.

Aequorin Bioluminescence Ca2+ Assays Aequorin assays were developed according to the methods of Button and Brownstein (39) and Stables et al. (40). The respective receptor-expressing HEK-293 cells stored at –80°C were thawed quickly at 37°C in a water bath and were cultured for transfection with the apo-aequorin gene construct. pcDNA3-cytAEQ or -mtAEQ was transiently transfected into HEK-BmDopR1–3 or -BmETHR cells using Lipofectamine as described above. The next day, the cells were washed twice with 1 ml of D-PBS and cultured in fresh culture medium for 1 day. After washing again with D-PBS, 1 ml of DMEM containing 0.1% FBS, 5 μM coelenterazine, and 30 μM reduced glutathione (DMEM : FBS : 5 mM methanolic coelenterazine solution with 30 μM glutathione : 30 mM aqueous glutathione solution = 1 ml : 1 μl : 1 μl : 1 μl) was added to the cells on a 35-mm dish. After incubation for 3–4 h at 37°C and with 5% CO2 followed by rinsing of the cells with 2 ï× 1 ml of D-PBS pre-warmed at 37°C, 1 ml of pre-warmed D-PBS with 10 mM EDTA was added to the cells. After incubation for 10 min in the CO2 incubator, the detached cells were collected with 2 × 1 ml of D-PBS in a 15-ml centrifuge tube. After centrifugation at 1,500 rpm for 2 min, the pelleted cells were gently suspended in 2.5 ml of pre-warmed extracellular buffer (140 mM NaCl, 20 mM KCl, 20 mM HEPES, 5 mM glucose, 1 mM MgCl2, 1 mM CaCl2, and 0.1 mg/ml bovine serum albumin, pH 7.4) per dish. The 15-ml tube containing the cell suspension was covered with aluminum foil and maintained at 37°C in a water bath before use. A 180-μl portion of the cell suspension was quickly added in a 1.5-ml microtube filled with a 20-μl aliquot of ligand solution in advance. Immediately after addition of the cell suspension, luminescence of the sample was counted for 30 s with a luminometer Lumi Counter NU-700 (Microtec, Chiba, Japan). The above procedure was repeated for each sample. EC50 values were estimated by the Probit method. To prepare the ligand solutions, test compounds were first dissolved and diluted in extracellular buffer or DMSO. When DMSO was used as a solvent, the final concentration was 1%. DMSO had no effect on the current aequorin assay. In agonist experiments in HEK-BmDopR cells (mainly HEK-BmDopR2 cells), biogenic amines (DA, OA, TA, and 5-HT), the DA receptor agonist 6,7-ADTN, and OA receptor agonists (DMCDM and NC-5) were used. In HEK-BmETHR cells, BmETH was tested as a putative endogenous peptide hormone. In antagonist experiments in BmDopR2 cells, increasing concentrations of Flu as the DA receptor antagonist, DMCDM, or NC-5 were premixed and tested with 10–8 M DA. 121

Acknowledgments We would like to thank Drs. Teiichi Tanimura and Hiroshi Ishimoto (Kyushu University, Japan) for kindly providing the plasmid vector containing an apo-aequorin cDNA. We greatly thank Dr. Yoshihisa Ozoe (Shimane University, Japan) for kindly providing DMCDM and NC-5. We also thank Dr. Honoo Satake (Suntory Foundation for Life Sciences) for kindly providing BmETH. This work was supported in part by KAKENHI (Grants-in-Aid for Scientific Research (C) (HO)). We would like to thank Editage (www.editage.jp) for English language editing.

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Chapter 7

Molecular Pharmacology and Physiology of Insect Biogenic Amine Receptors Jia Huang* Institute of Insect Sciences, Zhejiang University, Hangzhou 310058, China *E-mail: [email protected].

Insect biogenic amines play important roles in mediating behavioral and physiological processes as neurotransmitters, neuromodulators and neurohormones. They activate specific G-protein coupled receptors on the cell surfaces to induce downstream signaling pathways. These biogenic amine receptors show different pharmacological properties from their vertebrate counterparts and therefore become potential targets for pest control. However, their physiological functions are not well understood. We summarize our recent progress on insect biogenic amine receptors, and emphasize the characterizations of novel receptors and their functions in behavior and immunity.

Introduction In vertebrates, biogenic amines such as dopamine, serotonin, epinephrine and norepinephrine act physiologically as neurotransmitters, neuromodulators and neurohormones to regulate many important processes. In contrast, norepinephrine and epinephrine do not appear to be present in insects, as insects lack dopamine β-hydroxylase that converts dopamine to norepinephrine. In fact, their role is fulfilled by their invertebrate counterparts, the monoamines tyramine and octopamine (Figure 1). The insect biogenic amines carry out many of the physiological roles such as reproduction, development, growth, circadian rhythms, endocrine secretion, and behaviors. They exert their effects by binding to specific receptor proteins that belong to the superfamily of G-protein coupled receptors (GPCRs), many of which have been characterized not only from the fruit fly Drosophila melanogaster but also from several other insect species. Thus, blocking or over stimulating these GPCRs in insect pests may either © 2017 American Chemical Society

result in the death or reduce fitness to control pest populations. Therefore, insect biogenic amine receptors are potential targets for the development of next generation of insecticides. We used a combination of approaches from cell biology, molecular biology, neurobiology, immunology and genetics to study the pharmacology and physiology of biogenic amine receptors in lepidopteran pests and D. melanogaster. In the meantime, we also discovered several novel families of biogenic amine receptors.

Figure 1. Biosynthetic pathway of biogenic amines. (A) Biosynthetic pathway of dopamine, norepinephrine, tyramine and octopamine. DβH, dopamine β-hydroxylase; DDC, Dopa decarboxylase; TDC, tyrosine decarboxylase; TβH, tyramine β-hydroxylase; TH, tyrosine hydroxylase. (B) Serotonin biosynthetic pathway. Tryptophan hydroxylase (TRH/TPH), aromatic L-amino acid decarboxylase (AADC).

A Novel Octopamine Receptor Octopamine regulates many physiological processes in insects through specific octopamine receptors on the cell membranes. The first insect octopamine receptor gene oamb was isolated from the mushroom body of the fruit fly D. melanogaster (1). Subsequently, a variety of octopamine receptors were cloned from several other insect species. Based on the structural and signaling similarities between cloned D. melanogaster octopaminergic receptors and vertebrate adrenergic receptors. Evans and Maqueira (2) proposed a new classification system, which divided insect octopaminergic receptors into two classes: α-adrenergic-like receptors (OA1) and β-adrenergic-like receptors (OA2). Activation of OA1 receptors primarily leads to the elevation of [Ca2+]i 128

when expressed in cell lines (3, 4). The OA2 receptors are divided into three subclasses, which all increase intracellular cAMP levels (5). Both OA1 and OA2 receptors show high specificity to octopamine over other biogenic amines such as tyramine and dopamine.

Figure 2. Agonists and antagonists of insect biogenic amine receptors (listed in the order that they appear in the text). 129

We have cloned and functionally characterized an octopamine receptor from the rice stem borer, Chilo suppressalis, which is one of the most economically important rice pests in Asia, northern Africa, and southern Europe. Orthologous receptors have not been isolated from other invertebrate species. Structural and pharmacological studies demonstrated that this gene belongs to a novel family of octopamine receptors, which we named CsOA3 receptor (6). The CsOA3 encodes two polypeptides, CsOA3S and CsOA3L, that are generated by alternative splicing. CsOA3L differs from CsOA3S by the presence of an additional 30 amino acids within the third intracellular loop. Phylogenetic analysis clearly indicates that CsOA3 clusters with predicted orthologous genes of D. melanogaster and Tribolium castaneum in a distinct clade. A closely related clade contains the human α2-adrenergic receptors. When heterologously expressed, activation of CsOA3 by octopamine primarily leads to decrease forskolin-stimulated [cAMP]i. It can also be activated by tyramine, but with lower potency and efficacy. CsOA3 shows a distinct pharmacology to reported octopamine and tyramine receptors (7). Naphazoline and clonidine act as potent agonists (Figure 2). Phentolamine and epinastine are able to block CsOA3 while yohimbine, chlorpromazine and mianserin are not effective.

A Novel Serotonin Receptor Serotonin (5-hydroxytryptamine; 5-HT) is a small molecule found in organisms across the animal kingdom. It modulates a wide variety of processes in most vertebrates and invertebrates. Except the 5-HT3 receptor, a ligand-gated cation channel, vertebrate serotonin receptors have been classified into six main classes (5-HT1A/B/D/E/F, 5-HT2A/B/C, 5-HT4, 5-HT5A/B, 5-HT6, 5-HT7) and three of them are also found in insects (5-HT1A/B, 5-HT2A/B and 5-HT7). The 5-HT1 receptors couple preferentially to Gi/o proteins and inhibit cAMP production. The 5-HT2 receptors couple preferentially to Gq/11 proteins, which lead to an increase in [Ca2+]i. The 5-HT7 receptors couple preferentially to Gs proteins and induce cAMP production (8). We isolated a GPCR cDNA encoding a potential serotonin receptor from the caterpillar of the small white butterfly, Pieris rapae, which shares relatively low similarity to the known serotonin receptor families (9). After heterologous expression in HEK-293 cells, this new receptor can be activated by serotonin, but not other biogenic amines, in a concentration-dependent manner. We propose this receptor represents a new group of serotonin receptors, and designate it Pr5-HT8. Bioinformatic studies indicate that species orthologues of Pr5-HT8 are mainly found in lepidopteran and coleopteran pests such as the diamondback moth Plutella xylostella, the red flour beetle Tribolium castaneum and also in the genome of the malaria mosquito Anopheles gambiae. However, it is not found in the genome of honey bee Apis mellifera, parasitoid wasp Nasonia vitripennis or other mammals. The phylogenetic analysis also show that Pr5-HT8 receptor does not cluster with reported 5-HT receptors, but it clusters with the predicted insect 5-HT8 family in a distinct clade. Besides serotonin, classical serotoninergic agonists 5-methoxytryptamin, 8-OH-DPAT and 5-carboxamidotryptamine 130

(Figure 2) activate Pr5-HT8 to induce calcium response, suggesting that Pr5-HT8 selectively couples to Gq protein to activate phospholipase C, leading to a signaling cascade that ends with an elevation of [Ca2+]i. A surprising discovery is that methiothepin, a nonselective serotonin receptor antagonist, can also activate Pr5-HT8. However, SB 269970, SB 216641 and RS 127445 have no blocking effect except WAY 100635, a 5-HT1A antagonist, can inhibit serotonin-induced [Ca2+]i increases. Thus, the pharmacological profiles of Pr5-HT8 differ from that of other serotonin receptors.

Characterization of Three Dopamine Receptors Dopamine acts as an important neurotransmitter and neuromodulator to regulate a variety of physiological responses and behaviors in insects, such as learning and memory, cognition, sexual orientation, locomotion, phase change, etc. In vertebrates, five distinct dopamine receptors mediate all known functions of dopamine. They can be divided into two subfamilies based on their structural and pharmacological properties: D1-like receptors (D1 and D5), and D2-like receptors (D2, D3 and D4). Insect D1-like receptors also have two subtypes, DOP1 and DOP2, which elevate intracellular cAMP levels upon activation. However, DOP2 can also increases [Ca2+]i levels and is considered as a invertebrate-specific dopamine receptors which show similarities in structural, signaling and pharmacological properties with OA1 receptors. DOP3 are functionally D2-like and inhibit adenylyl cyclase (10). A Drosophila GPCR (DopEcR) that can be activated by both DA and ecdysteroids has been reported and it shares a high level of amino-acid sequence similarity with β-adrenergic receptors. We have pharmacologically characterized three types of dopamine receptors, CsDOP1, CsDOP2 and CsDOP3, from the rice striped stem borer, Chilo suppressalis (10). They all show considerable sequence similarity with orthologous dopamine receptors and phylogenetic analysis also clusters the receptors within their each group. Transcript levels of CsDOP1, CsDOP2 and CsDOP3 are all significantly high in the central nervous system, indicating their important roles in neural processes. After heterologous expression in HEK 293 cells, CsDOP1, CsDOP2 and CsDOP3 are dose-dependently activated by dopamine and synthetic dopaminergic agonists (Figure 2). The rank orders of agonist activity for CsDOP1 is dopamine > bromocriptine > 6,7-ADTN > pramipexole. For CsDOP2, only dopamine and 6,7-ADTN can activate the receptor significantly and bromocriptine also shows high agonist activity on CsDOP3. The rank order of potency of tested antagonists on CsDOP1 is the following: butaclamol ≥ SCH-23390 > chlorpromazine ≥ flupenthixol > mianserin > phentolamine ≥ spiperone > yohimbine > propranolol ≥ ketanserin. The rank order for CsDOP2 is: chlorpromazine ≥ clozapine ≥ SCH-23390 ≥ epinastine ≥ mianserin ≥ flupenthixol ≥ butaclamol > phentolamine > prazosin > spiperone ≥ sulpiride. For CsDOP3, the rank order of potency of antagonists is the following: epinastine > mianserin > SCH-23390 > chlorpromazine. 131

Tyramine Receptor Modulates Courtship Behavior In insects, tyramine is a decarboxylated product of tyrosine and further hydroxylated to produce octopamine. For a long time, it was assumed to serve as a biosynthetic precursor of octopamine, rather than as a neuroactive substance. Therefore, not much is known about the physiological role of tyramine. In the last decade, increasing reports have supported the hypothesis that tyramine is not only a precursor but also a potential genuine signaling molecule in a variety of physiological processes. However, it is still unclear whether tyramine acts as an independent neuromodulator in vivo due to the lack of solid genetic evidences. Although there is one point-mutation allele for Drosophila Tdc2 (Tdc2RO54), which is the major tyrosine decarboxylase in central nervous system. Thus, the Tdc2RO54 allele abolishes most tyramine and octopamine; and the Tβh null allele Tβhnm18 allele has no octopamine but 10 times higher of tyramine compared to control (11). Considering even simple behaviors may include complex neuro-circuits with multiply neurotransmitters and neuromodulators involved, it is impossible to make clear conclusions about the role of tyramine with above two alleles. Three tyramine receptors were identified in Drosophila: Oct-TyrR (CG7485), TyrR (CG7431) and TyrR2 (CG16766). Oct-TyrR couples to Gi and Gq proteins when heterologously expressed in cell lines and shows a pharmacology profile with slightly higher potency to tyramine over octopamine and dopamine (12, 13). TyrR2 is also like Oct-TyrR to coupled to Gi and Gq proteins but have a higher preference for tyramine than other amine (14). However, TyrR is an unusual insect biogenic amine receptor since it shows a very high specificity for tyramine (EC50 = 3 x 10-7 M). Structurally related biogenic amines, such as octopamine, synephrine and dopamine did not show any effects up to a concentration of 100 μM on TyrR (15) or on its species homologue from the silkworm, Bombyx mori (16). Nevertheless, of all the octopamine and tyramine GPCRs, TyrR is the only one to show activity to tyramine but not other biogenic amines. Therefore, we reasoned that we can disclose the physiological role of tyramine through generation of a TyrR knock-out allele in that any defect of TyrR mutant can only be attribute to its solo endogenous ligand, tyramine (17). To simultaneously generate mutations and gene reporters, we used ends-out homologous recombination to insert the GAL4 gene at the site of the normal TyrR translation initiation codon. At the same time, we deleted a ~0.7 kb exon including the N-terminal coding region and the first two transmembrane domains of TyrR. These TyrRGal4 males show strong male-male courtship behaviors (Figure 3). When 8-10 TyrRGal4 males are introduced into a small Petri dish together, they serially chase each other to exhibit a chain formation in which males court each other and produce courtship song. Either knocking down TyrR expression by RNAi or activation of TyrR neurons also phenocopied the chaining behavior. These results demonstrate that tyramine might act as an inhibitory neuromodulator in vivo, which is consistent with previous in vitro physiological studies showing that tyramine reduces the amplitude of excitatory junction potential (EJP) in neuromuscular junctions. Genetic and behavioral studies further indicates that TyrR activity is required in a small group of neurons in the brain, which may form 132

synaptic connections with the male-specific Fruitless protein (FruM) neurons to regulate courtship (17).

Figure 3. TyrR Gal4 mutant showed enhanced courtship behavior. (A) The expression pattern of the TyrR reporter (TyrRGal4/+) in the brain. The red arrows indicate the location of some of the neurons expressing the TyrR reporter. SMP, superior medial protocerebrum; PLP, posteriorlateral protocerebrum; IPS, inferior posterior slope; GNG, gnathal ganglia. MB, mushroom body; AL, antennal lobe; OL, optical lobe. (B) Model illustrating the proposed role of TyrR and IPS neurons in the male fly brain in the regulation of courtship behaviors. Tyramine and octopamine are released from Tdc2 neurons in response to a close encounter with a male. Octopamine promotes male aggression. The tyramine activates the TyrR, which in turn inhibits TyrR-expressing IPS (TyrRIPS) neurons. In the absence of inhibition, the TyrRIPS neurons release acetylcholine, which promotes courtship. (Reproduced with permission from reference (17) Copyright 2016 Elsevier.).

Serotonin Receptors Regulate Immunity The immune system needs to be tightly regulated and highly responsive to changes in external and internal environments. Neurohormones affect the function of the immune system, but how they do so is not well understood. Serotonin modulates both neural and immune responses in mammals, where various types of immune cells that engulf foreign particles or microorganisms through a process called phagocytosis, have receptors for serotonin on their cell surface and are 133

activated when serotonin is present. Serotonin is also known to influence many processes in insects, such as appetite, sleep and reproduction, but its role in insect immunity is still unclear. Besides, it is also elusive how the immune response is regulated and coordinated by neurohormones at the level of the whole organism.

Figure 4. A schematic diagram of serotonin signaling on hemocyte phagocytosis. Lipopolysaccharide (LPS) enhances the expression of tryptophan hydroxylase (TPH), which catalyzes tryptophan into serotonin via 5-hydroxy tryptophan (5-HTP). Serotonin, which is secreted from hemocytes, activates the hemocyte-membrane receptor 5-HT1B and 5-HT2B. The immune responses of P. rapae are labeled in purple: activation of 5-HT1B promotes hemocyte phagocytosis and activation of 5-HT2B lead to opposite effects. LPS increases 5-HT1B expression but decreases that of 5-HT2B. The immune responses of Drosophila are labeled in green arrows: activation of 5-HT1B promotes hemocyte phagocytosis and activation of 5-HT2B leads to the same effects. (Reproduced from reference (18). Permission is not required from eLife). We found that hemocytes (insect blood cells) in the caterpillar, Pieris rapae are able to synthesize and release serotonin following activation by lipopolysaccharide (LPS), the major component of the outer membrane of Gram-negative bacteria. The inhibition of a serotonin-generating enzyme with either pharmacological blockade or siRNA silencing results in significantly decreased hemocyte phagocytosis. Two distinct serotonin receptors (5-HT1B and 5-HT2B) are expressed on the cell surfaces of naive hemocytes. Using selective antagonists and RNAi, we found that inhibition of 5-HT1B decreases hemocyte phagocytosis. However, inhibition of 5-HT2B enhances hemocyte phagocytosis. Interestingly, 5-HT1B is dramatically up-regulated following hemocyte activation, but 5-HT2B is significantly down-regulated (Figure 4). We confirmed the role of 5-HT receptors in vivo using the D. melanogaster. D. melanogaster 5-HT1B 134

deficient flies were more vulnerable to bacterial infections due to their poor phagocytosis ability, indicating that the 5-HT1B-mediated hemocyte phagocytosis is critical in the insect immune response to invading organisms. Flies expressing 5-HT1B or 5-HT2B RNAi in hemocytes also showed similar sensitivity to infection. This is the first molecular demonstration of serotonin receptors in an insect immune cell. This result establishes, at the molecular level, that animals from a phylum outside of the vertebrates could also use serotonin and its receptors to connect the nerve system to immune function. To our knowledge, it is also the first genetic evidence to show that loss of a neurohormone receptor leads to immunodeficiency. Further studies are needed to examine how physiological changes such as behaviors, nutritional state, and stress will affect insect immunity through serotonin signaling pathways (18).

Biogenic Amine Receptors as Pest Control Targets Many insecticidal chemicals can act on insect biogenic amine receptors (Figure 5). The octopamine receptor gets more attention since it is a bona fide target of a class of commercial insecticides, the formamidines. In 1980, Hollingworth (19) and Evans (20) first reported that the formamidine acaricide/insecticide, chlordimeform (CDM) and its N-demethylated derivative (DMCDM) acted as octopamine receptor agonists by physiological studies using insect native organs or tissues. We confirmed that DMCDM can activate the Bombyx mori α-adrenergic-like octopamine receptor (BmOAR1) to induce both cAMP and Ca2+ responses when expressed in HEK-293 cell lines (21). We further found that CDM and another formamidine insecticide, amitraz can also directly activate the Chilo suppressalis β-adrenergic-like octopamine receptor (CsOA2B2) and the above novel octopamine receptor CsOA3 (unpublished data). Recently, Kita et al. reported that amitraz and its metabolite N2-(2,4-dimethylphenyl)-N1-methyformamidine (DPMF) potently activate Bombyx mori α- and β-adrenergic-like octopamine receptors but their potencies on these two receptors are quite different (22). Besides, insecticidal essential oils are also believed to act on octopamine receptors (23). Enan found that α-adrenergic-like octopamine receptors from D. melanogaster and Periplaneta americana respond to cinnamic alcohol, eugenol and trans-anethole (24). However, we found that only eugenol at 100 μM, but not cinnamic alcohol and trans-anethole, can activate BmOAR1 to induce Ca2+ response (21). Tyramine receptors can also be activated by CDM (7) and plant essential oils (25, 26). For dopamine receptors, a yellow fever mosquito (Aedes aegypti) dopamine receptor (AaDOP1) was used in a chemical library screen, amitriptyline and doxepin were identified as antagonists and showed high toxicity in subsequent A. aegypti larval bioassays (27). There are few papers about insecticidal compounds that target serotonin receptors. However, one study used the 5-HT1A agonist PAPP (1-[(4-aminophenyl)ethyl]-4-[3-(trifluoromethyl)phenyl]piperazine) as a lead compound for new insecticides design and they found that most synthesized PAPP derivatives displayed certain growth-inhibiting activities or larvicidal activities against the armyworm, Pseudaletia separate (28). 135

Figure 5. Insecticidal chemicals targeting insect biogenic amine receptors (listed in the order that they appear in the text).

Conclusion Insect biogenic amine receptors have unique pharmacological profiles, which are different from their counterparts in mammals. Some of them, like 5-HT8 receptor, are even restricted in few insect species (9). They are also involved in regulating a series of behaviors and key physiological functions. While octopamine receptors are an established insecticide target, there is no commercial insecticide targeting other biogenic amine receptors. Thus, insect biogenic amine receptors merit more studies which will promote the development of novel insecticides with novel mode of actions.

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tyramine receptor selectively coupled to intracellular calcium mobilization. Insect. Biochem. Mol. Biol. 2009, 39, 842–849. Huang, J.; Liu, W.; Qi, Y. X.; Luo, J.; Montell, C. Neuromodulation of courtship drive through tyramine-responsive neurons in the Drosophila brain. Curr. Biol. 2016, 26, 2246–2256. Qi, Y. X.; Huang, J.; Li, M. Q.; Wu, Y. S.; Xia, R. Y.; Ye, G. Y. Serotonin modulates insect hemocyte phagocytosis via two different serotonin receptors. eLife 2016, 4, e04805. Hollingworth, R. M.; Murdock, L. L. Formamidine pesticides: octopaminelike actions in a firefly. Science 1980, 208, 74–76. Evans, P. D.; Gee, J. D. Action of formamidine pesticides on octopamine receptors. Nature 1980, 287, 60–62. Huang, J.; Hamasaki, T.; Ozoe, Y. Pharmacological characterization of a Bombyx mori α-adrenergic-like octopamine receptor stably expressed in a mammalian cell line. Arch. Insect Biochem. Physiol. 2010, 73, 74–86. Kita, T.; Hayashi, T.; Ohtani, T.; Takao, H.; Takasu, H.; Liu, G.; Ohta, H.; Ozoe, F.; Ozoe, Y. Amitraz and its metabolite differentially activate α- and βadrenergic-like octopamine receptors. Pest Manage. Sci. 2017, 73, 984–990. Enan, E. Insecticidal activity of essential oils: octopaminergic sites of action. Comp. Biochem. Physiol., Part C: Toxicol. Pharmacol. 2001, 130, 325–337. Enan, E. E. Molecular and pharmacological analysis of an octopamine receptor from American cockroach and fruit fly in response to plant essential oils. Arch. Insect Biochem. Physiol. 2005, 59, 161–171. Enan, E. E. Molecular response of Drosophila melanogaster tyramine receptor cascade to plant essential oils. Insect Biochem. Mol. Biol. Molec. 2005, 35, 309–321. Gross, A. D.; Temeyer, K. B.; Day, T. A.; Perez de Leon, A. A.; Kimber, M. J.; Coats, J. R. Interaction of plant essential oil terpenoids with the southern cattle tick tyramine receptor: A potential biopesticide target. Chem. Biol. Interact. 2017, 263, 1–6. Meyer, J. M.; Ejendal, K. F.; Avramova, L. V.; Garland-Kuntz, E. E.; GiraldoCalderon, G. I.; Brust, T. F.; Watts, V. J.; Hill, C. A. A "genome-to-lead" approach for insecticide discovery: pharmacological characterization and screening of Aedes aegypti D(1)-like dopamine receptors. PLoS Neglected Trop. Dis. 2012, 6, e1478. Cai, M.; Li, Z.; Fan, F.; Huang, Q.; Shao, X.; Song, G. Design and synthesis of novel insecticides based on the serotonergic ligand 1-[(4-aminophenyl)ethyl]-4-[3-(trifluoromethyl)phenyl]piperazine (PAPP). J. Agric. Food Chem. 2010, 58, 2624–2629.

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Chapter 8

A Review of Aminothiazoline Chemistry Barbara Wedel,*,1 Wolfgang von Deyn,2 Sebastian Soergel,2 Matthias Pohlman,1 Douglas Anspaugh,1 Ramani Kandasamy,1 Fae Malone,1 Daniel Houtz,1 Nancy Rankl,1 John Dorsch,1 Lynn Stam,1 Brecht London,1 Ronan le Vezouet,2 Christopher Koradin,2 and Markus Kordes2 1BASF

Corp., 26 Davis Drive, Research Triangle Park, North Carolina 27709-3528, United States 2BASF SE, B009, Ludwigshafen, Delaware 67056, United States *E-mail: [email protected].

Aminothiazoline (ATZ) chemistry was developed starting from a random in vivo screening hit with moderate activity in aphids. A series of compounds was synthesized to increase potency and breadth of spectrum towards other piercing sucking insects. ATZs showed no cross-resistance to chloronicotinyl insecticides (CNIs) and pyrethroids and controlled multi-resistant whitefly species. The main symptoms in aphids were wandering behavior along with fast feeding cessation, the speed of control was slow. As to the mode of action, based on structural similarity of the original screening hit with a known octopaminergic agonist we closer examined the possibility of ATZs signaling through insect octopamine receptors. Although in vitro activity in the micromolar range could be observed with ATZs on some octopamine receptors, there was no clear in vivo-in vitro correlation suggesting a possibly new mode of action for this chemistry. Initial Tox- and Ecotox profiles were very positive, but late stage toxicity testing raised concern that together with a lack of systemicity and limited spectrum prompted us to stop further activity on ATZs.

© 2017 American Chemical Society

Discovery and Overview of Aminothiazolines In this book chapter aminothiazoline chemistry is reviewed, which provides excellent control of piercing-sucking insects. The initial discovery, structure-activity relationship, mode of action work and biological spectrum will be covered in the following sections. The chemistry was developed from a random screening hit, the N-(1,2-diphenylethyl)aminooxazoline derivatives shown in Figure 1, which showed moderate activity on cotton aphid (Aphis gossypii) and green peach aphid (Myzus persicae). Chemical variation of the scaffold resulted in increased activity on aphids and expansion of the spectrum to whiteflies. Further structure optimization led to the lead analog shown on the right in Figure 1, which showed excellent activity on aphids and whiteflies and further expansion of the spectrum to thrips and hoppers. Using a scaffold hopping approach, Lambert et al. recently reported insecticidal activity for structurally related thioureas and isothioureas, that controlled Myzus persicae and Bemisia tabaci in laboratory bioassays (1).

Figure 1. Development of aminothiazoline chemistry from the original Aminooxazoline screening hit into a screening hit cluster with heteroatom (X), prodrug group (PG) and R-group variation on both phenyl rings (R1-R6). The lead Aminothoiazoline analog with an extended pest spectrum is shown on the right side.

The mode of action appears to be novel and will be discussed in more detail below. The route of exposure was mainly contact with limited oral activity. While the speed of aphid control was slow, feeding cessation was fast, which is an important factor in virus transmission. No cross-resistance to conventional insecticide classes such as CNIs and pyrethroids was observed. Translaminar plant movement could be seen for several aminothiazoline analogs, but no appreciable systemicity could be detected. 140

The regulatory profile of the lead aminothiazoline analog was very favorable: acute toxicity in rats was low (LD50 > 500 mg/kg) with no observed eye or skin irritation and a negative Ames test (non-mutagenic). With regards to ecotoxicology, there were no major concerns for aquatic organisms (acute fish, Daphnia and Chironomus) and low acute toxicity for wildlife (quail) and bees. Environmental fate studies showed a high organic carbon-water partition co-efficient and therefore no leaching risk. Soil degradation was moderate to slow and the compound was rapidly degraded in air.

Figure 2. Summary of structure-activity relationship of aminothiazoline chemistry. Heterocycle substitutions (1.) including prodrug groups (PG), A-ring (2.) and B-ring substitutions (3.) providing best in vivo activity are depicted.

Aminothiazoline Chemistry Key structure-in vivo activity relationships are summarized in Figure 2. 5-ring heterocycles showed best activity with thiazolines showing higher activity than oxazolines or imidazolines (Figure 2). While heterocyclic substitutions on the B-ring were tolerated, but showed weakened activity, heterocyclic A-ring substitutions generally lost biological activity with the exception of thiophene. Figure 3 shows the preferred synthesis route for aminothiazolines: Commercially available 2,3-dimethylbenzaldehyde was treated with lithium bis(trimethylsilyl)amide to form a silyl-imine. A Grignard reagent – prepared from the respective chloride – was added afterwards yielding the diphenylethylamine. With the addition of thiophosgene an isothiocyanate was produced in very good yields. Ethanolamine was added to form a thiourea compound which was subsequently dehydrated under various conditions to form the lead aminothiazoline. 141

Figure 3. Synthesis route for the lead aminothiazoline analog. The synthesis route was very robust and reproducible from small mg to kg amount scale.

Aminothiazoline Mode of Action Octopamine is an essential neurotransmitter, neuromodulator and neurohormone in the insect nervous sytem involved in coordination of locomotor behavior, modulation of sensory responses, learning and memory as reviewed elsewhere (2, 3). Octopamine and tyramine receptors are interesting targets for the development of Insecticides and octopamine receptor agonists such as amitraz (AM) and chlordimeform (CDM) have long been know to have insecticidal/acaricidal properties (4, 5). Both formamidines are thought to be pro-insecticides, which are metabolized within the insect to the active metabolites BTS27271 and demethylchlordimeform (DCDM), respectively (Figure 4). Searching for new classes of octopamine agonists, Jennings et al. discovered that some 2-aminooxazolines identified in an in vitro approach were aphicidal and acaricidal, and exhibited symptomology characteristic for octopaminergic agonists (6). The phenylaminooxazoline AC-6, one of the most potent compounds in the series, showed structural similarity to our initial screening hit, which had also been an aminooxazoline (Figure 4). This prompted us to investigate octopamine receptor activation as a putative mode of action for our aminothiazoline chemistry. We decided to develop a cell line stably expressing the Drosophila melanogaster octopamine receptor Oa2 (Dm-Oa2), which is also known in literature as OctR or Octβ1R and corresponds to CG6919. Insect neurohormone GPCRs and octopamine receptors in particular have been reviewed elsewhere (7, 8). A classification system based on structural similarities and signaling properties with vertebrate adrenergic receptors was proposed in 2005 by Evans 142

and Maqueira (9). They suggested to distinguish between α-adrenergic-like octopamine receptors (OctαRs), β-adrenergic-like octopamine receptors (OctβRs) and tyraminergic receptors. Based on literature evidence, Dm-Oa2 activation causes elevation of cAMP levels when expressed in HEK293 cells (10). We stably expressed Dm-Oa2 in a CHO-Gα16 background thus forcing the coupling of the GPCR to increases in cytosolic calcium levels via stimulation of PLC-β as shown in Figure 5.

Figure 4. Structures of the initial in vivo screening hit, the octopaminergic agonist AC-6 described in(6), amitraz and its active metabolite BTS27271 and chlordimeform and its active metabolite DCDM. BTS27271 is also known in literature as DPMF (N-(2,4-dimethylphenyl)-N‘-methylformamidine).

We pharmacologically characterized the Dm-Oa2/CHO-Gα16 cell line as summarized in Table 1, using the parental CHO-Gα16 line as a control. The natural ligand octopamine activated Dm-Oa2 with an EC50 of 13.6 nM. Agonists and antagonists of mammalian GPCRs described in literature to affect insect octopamine receptors were also tested. Mianserin and cyproheptadine acted as antagonists of Oa2 with an IC50 of 31.2 nM and 50.8 nM, respectively. Naphazoline, tolazoline and clonidine had agonist activity on Dm-Oa2 with a rank order of potency of naphazoline (5.7 nM) > clonidine (75.1 nM) > tolazoline (217.5 nM). As to be expected, the pro-insecticide forms of the two formamidines amitraz (EC50 = 176 nM) and CDM (EC50 = 13600 nM) were less potent on Dm-Oa2 than the respective active metabolites BTS27271 and DCDM (EC50 of 12 nM and 16 nM, respectively; see Table 1). 143

Figure 5. Dm-Oa2 was stably expressed in a CHO-Ga16 background linking GPCR activation to increases in intracellular Ca2+ levels via activation of PLC and activation of the ER-residing IP3 receptor. The increased calcium release can be measured with fluorescent calcium indicator dyes such as Fluo-4.

As shown in Table 1, the lead ATZ analog did not exhibit high potency on Dm-Oa2 with an EC50 only in the micromolar range. We also tested the lead ATZ in antagonist mode challenging the receptor with 100nM Octopamine, but could not observe any inhibitory effect. We were wondering whether species differences might be responsible for the lack of activity of the lead on Dm-Oa2 and therefore also cloned and stably expressed the corresponding aphid receptor Mp-Oa2 (Myzus persicae) in the same CHO-Gα16 background as the Drosophila receptor Dm-Oa2. However, in both cell lines the lead ATZ analog was 850-fold (Dm-Oa2) and 80fold less potent (Mp-Oa2) than the active form of amitraz, BTS27271. We tested an entire series of ATZ derivatives on the two Oa2-receptor expressing cell lines, which exhibited a range of potencies, but no clear in vitro - in vivo correlation was observed (data not shown). 144

Table 1. Pharmacological Characterization of the Dm-Oa2/CHO-Gα16 Cell Line. Agonist or Antagonist Activity of Modulators of Mammalian GPCRs, Formamidines, Octopamine and the Lead ATZ Compound Were Determined.

We next decided to establish an octopamine receptor panel. We included the β-adrenergic-like octopamine receptors (Dm-Oa2, Mp-Oa2, Dm-Octβ2R Tc-Octβ3R, the α-adrenergic-like octopamine receptor Dm-oamb, and the tyraminergic receptor Dm-Tyr, also referred to as Oct-TyrR1, a Type 1 receptor (Table 2). The receptors were cloned from Drosophila melanogaster (Dm), Myzus persicae (Mp) or Tribolium confusum (Tc) as indicated and stably expressed in a CHO-Gα16 background as described above. As shown in Table 2, octopamine exhibited highest potency on Tc-Octβ3R with an EC50 of 2 nM, while Dm-TyrR showed clear preference for tyramine over octopamine (EC50 of 165 nM; tyramine). In the cell lines tested (Dm-Oa2, Mp-Oa2 and Dm-OAMB), formamidine pro-insecticides showed lower potency than their corresponding active forms (Table 2). Interestingly, Kita et al. recently reported that the active amitraz metabolite DPMF (BTS27271) was more potent in increasing intracellular cAMP levels signaling through b-adrenergic-like octopamine receptors (EC50 = 79.6 pM) than elevating intracellular Ca2+ levels signaling through α-adrenergic-like octopamine receptors EC50 = 1.17 nM (11). Since we forced coupling through Gα16 and therefore a calcium readout in our assay panel, 145

we may not have been able to see this differential activation in our system. The above mentioned phenylaminooxazoline AC-6 that showed structural similarity with our initial in vivo screening hit, exhibited nM potency on Dm-Oa2 and Mp-Oa2. However, our ATZ lead analog exhibited only µM potency on Dm-Oa2, Mp-Oa2, Tc-Octβ3R and Dm-oamb while being inactive on Dm-Octβ2R and Dm-TyrR. There was no antagonist activity for the ATZ lead on any of tested receptors when challenged with 100nM octopamine or 250nM tyramine, respectively.

Table 2. Agonist Potencies of Octopamine, Formamidines, AC-6 and the ATZ lead on an Octopamine Receptor Panel. An Asterisk* Indicates That an EC50 Could Not Be Determined since No Detectable Increase in Intracellular Calcium Could Be Detected.

With respect to symptomology, chlordimeform-treated aphids showed hyperactivity while ATZ treated aphids exhibited mostly extensive wandering behavior. Our compiled data suggest a novel mode of action for ATZ, different from existing octopamine receptor agonists. Given that there is micromolar activity on some of the octopamine receptors one might speculate that ATZs might act on another insect GPCR. 146

Aminothiazoline Biology As mentioned above the original aminooxazoline screening hit shown in Figure 1 exhibited only moderate activity on the tested aphids (Aphis gossypii and Myzus persicae). With chemical variation, compounds with increased activity on aphids and widened spectrum to whiteflies could be identified. Further structure optimization yielded the ATZ lead analog, which showed excellent activity on aphids in the range of commercial standards, also controlled whiteflies including the difficult to manage Q biotype and additionally expanded the spectrum to thrips and hoppers as summarized in Table 3.

Table 3. Efficacy and Residual Activity of the Lead ATZ analog.

147

Summary and Conclusions ATZ chemistry was developed from a random in vivo screening hit and shows preference for controlling piercing-sucking insects. Although the speed of control was slow, feeding cessation was fast with aphids exhibiting a wandering behavior. In vitro activity on octopamine receptors was observed for several ATZ analogs, but there was no clear in vivo-in vitro correlation suggesting a possibly new mode of action for this chemistry. Attractive features of the chemistry were its lack of cross-resistance to CNIs and pyrethroids and very positive initial Tox- and Ecotox profiles. However late stage toxicity testing raised some concerns that together with a lack of systemicity and somewhat limited spectrum prompted us to stop activity on this chemistry.

References 1.

Lambert, W. T.; Goldsmith, M. E.; Sparks, T. C. Insecticidal activity of novel thioureas and isothioureas. Pest Manage. Sci. 2017, 73, 743–751. 2. Roeder, T. Tyramine and octopamine: ruling behavior and metabolism. Ann. Rev. Entomol. 2005, 50, 447–477. 3. Roeder, T. Octopamine in invertebrates. Prog. Neurobiol. 1999, 59, 533–61. 4. Hirashima, A. Tyramine and octopamine receptors as a source of biorational insecticides. In Biorational Control of Arthropod Pests: Application and Resistance Management; Ishaaya, I., Horowitz, A. R., Eds.; Springer Netherlands: Dordrecht, 2009; pp 83−109. 5. Hollingworth, R. M. Chemistry, biological activity, and uses of formamidine pesticides. Environ. Health Perspect. 1976, 14, 57–69. 6. Jennings, K. R.; Kuhn, D. G.; Kukel, C. F.; Trotto, S. H.; Whitney, W. K. A biorationally synthesized octopaminergic insecticide: 2-(4-chloro-otoluidino)-2-oxazoline. Pestic. Biochem. Physiol. 1988, 30, 190–197. 7. Hauser, F.; Cazzamali, G.; Williamson, M.; Blenau, W.; Grimmelikhuijzen, C. J. A review of neurohormone GPCRs present in the fruitfly Drosophila melanogaster and the honey bee Apis mellifera. Prog. Neurobiol. 2006, 80, 1–19. 8. Ohta, H.; Ozoe, Y. Chapter two - molecular signalling, pharmacology, and physiology of octopamine and tyramine receptors as potential insect pest control targets. In Advances in Insect Physiology; Ephraim, C., Ed.; Academic Press, 2014; Vol. 46, pp 73−166. 9. Evans, P. D.; Maqueira, B. Insect octopamine receptors: a new classification scheme based on studies of cloned Drosophila G-protein coupled receptors. Invert. Neurosci. 2005, 5, 111–118. 10. Balfanz, S.; Strunker, T.; Frings, S.; Baumann, A. A family of octopamine receptors that specifically induce cyclic AMP production or Ca2+ release in Drosophila melanogaster. J. Neurochem. 2005, 93, 440–451. 11. Kita, T.; Hayashi, T.; Ohtani, T.; Takao, H.; Takasu, H.; Liu, G.; Ohta, H.; Ozoe, F.; Ozoe, Y. Amitraz and its metabolite differentially activate α- and βadrenergic-like octopamine receptors. Pest Manage. Sci. 2017, 73, 984–990. 148

Editors’ Biographies Aaron Gross Dr. Aaron Gross is an Assistant Professor of Physiology and Toxicology in the Department of Entomology at Virginia Polytechnic Institute and State University. He earned his doctoral degree from Iowa State University of Science and Technology, under the supervision of Drs. Joel Coats and Michael Kimber. His postdoctoral research training was at the Emerging Pathogens Institute, University of Florida, under the supervision of Dr. Jeffrey Bloomquist. His research interest include the discovery, and understanding the biochemical and neurophysiological mechanisms of action of naturally occurring and synthetic pesticides, with the goal of controlling arthropods that are important vectors of human and animal health.

Yoshihisa Ozoe Dr. Yoshihisa Ozoe is a specially appointed professor at Shimane University, Japan. He earned his doctoral degree in agricultural chemistry from Kyushu University, Japan, in 1982 under the supervision of Professor Morifusa Eto. He joined Dr. Fumio Matsumura’s group at Michigan State University (1982-1984) and at University of California - Davis (1991). His research focus is on ligand-gated ion channels and G protein-coupled receptors as targets of insecticides. He is the recipient of the PSSJ High-Prospectiveness Award (1985), the PSSJ Prominent-Achievement Award (2004), and the ACS International Award for Research in Agrochemicals (2016).

Joel Coats Joel Coats is Distinguished Professor of Entomology & Toxicology at Iowa State University. He is an insect toxicologist with expertise in natural products as insecticides and insect repellents, including investigations of their selectivity, mechanisms of action, metabolism, synthesis of biorational derivatives and analogs, and quantitative structure-activity relationships (QSAR). A current focus is on uses of terpenes from plant essential oils as repellents, insecticides or synergists. Joel holds 9 patents, has published 11 books and over 200 scientific papers/review articles/book chapters; he started the Toxicology Graduate Program at Iowa State that has run continuously since 1985, and he currently serves as major professor for 6 Ph.D. students and has served as major professor for 45 previous graduate students. © 2017 American Chemical Society

Indexes

Author Index Akamatsu, M., 41 Anspaugh, D., 139 Asaoka, K., 109 Balfanz, S., 85 Baumann, A., 85 Blankenburg, S., 85 Blenau, W., 85 Coats, J., ix Dorsch, J., 139 Gross, A., ix Hill, C., 55 Houtz, D., 139 Huang, J., 127 Iwai, T., 41 Jones, A., 1 Kandasamy, R., 139 Koradin, C., 139 Kordes, M., 139 Kuroda, K., 41 le Vezouet, R., 139

London, B., 139 Malone, F., 139 Matsuda, K., 41 Mitsumasu, K., 109 Nagasaki, K., 41 Nakao, T., 23 Ohta, H., 109 Ozoe, Y., ix, 41 Pohlman, M., 139 Rankl, N., 139 Sakamoto, T., 41 Sharan, S., 55 Soergel, S., 139 Stam, L., 139 Tanaka, K., 41 Tanaka, Y., 109 Taylor-Wells, J., 1 von Deyn, W., 139 Wedel, B., 139 Yaginuma, T., 109

153

Subject Index A Aminothiazoline chemistry, review, 139 aminothiazoline biology, 147 lead ATZ analog, efficacy and residual activity, 147t aminothiazoline chemistry, 141 lead aminothiazoline analog, synthesis route, 142f aminothiazoline mode of action, 142 CHO-Ga16 background linking GPCR activation, Dm-Oa2, 144f Dm-Oa2/CHO-Gα16 cell line, pharmacological characterization, 145t initial in vivo screening hit, structures, 143f octopamine, agonist potencies, 146t aminothiazolines, discovery and overview, 140 aminothiazoline chemistry, development, 140f aminothiazoline chemistry, summary of structure-activity relationship, 141f

G GABA antagonist γ-benzene hexachloride, 41 discussion, 50 γ-BHC, docking, 51f introduction, 42 BHC isomers, conformations, 43f materials and methods activities, evaluation, 44 chemicals, 43 fluorometric imaging plate reader (FLIPR) membrane potential (FMP) assay, 45 [3H]4’-ethynyl-4-n-propylbicycloorthobenzoate ([3H]EBOB) binding assay, 44 insecticidal activity, 44 results Drosophila melanogaster GABA receptor, FLIPR membrane potential (FMP) assay, 50f

evaluation of in vitro GABA antagonistic activity, [3H]EBOB binding assay, 49 FLIPR membrane potential (FMP) assay, 49f FMP assay, 49 GABA antagonist, 46f insecticidal activities, 45 insecticidal activities against housefly and GABA antagonist activities, 47t insecticidal activity and poisoning symptom, 48t various isomers of BHC, HeptaCl and OctaCl, insecticidal activities against housefly, 48t GPCR-targeting insecticides, potential, 55 arthropod GPCRs, pharmacology, 61 Aedes aegypti AaDOP1 and AaDOP2 receptors, pharmacological characterization, 67f class A neurohormone GPCRs, 69 inhibition of dopamine-stimulated IP1 response, IC50 values, 68t invertebrate class A, summary, 63t vertebrate DARs, 65 conclusions, 74 GPCR-insecticides, challenges to development, 72 GPCR-targeting insecticides, argument, 60 G protein-coupled receptors (GPCRs), 58 G protein-coupling mechanisms, schematic diagram, 59f insecticide targets, development of arthropod GPCRs, 70 new insecticidal chemistries, drug discovery and development pipeline, 71f vector control, call for new mode of action insecticides, 56

I Insect biogenic amine receptors, molecular pharmacology and physiology courtship behavior, tyramine receptor, 132

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courtship behavior, TyrR Gal4 mutant, 133f introduction, 127 biogenic amines, biosynthetic pathway, 128f novel octopamine receptor, 128 insect biogenic amine receptors, agonists and antagonists, 129f novel serotonin receptor, 130 pest control targets, biogenic amine receptors, 135 insect biogenic amine receptors, insecticidal chemicals, 136f serotonin receptors regulate immunity, 133 serotonin signaling, schematic diagram, 134f three dopamine receptors, characterization, 131 Insect GABA receptor, variations, 1 alternative splicing on RDL receptor pharmacology, effects, 9 post-transcriptional modifications, schematic illustration, 10f conclusions and future prospects, 13 environmentally-induced variations, 4 A2’ mutation, resistance mutations found, 7t naturally-occurring variations, 9 RDL receptor pharmacology, effects of RNA editing, 11 Interspecies insect GABAB receptor heteromer, 85 discussion different GABAB receptors, pharmacological characteristics, 101t exchanging orthologous GB2 subunits, 101 interspecies PeaGB1/DmGB2 receptor heteromer, pharmacological properties, 99 PeaGB1/DmGB2 receptors, efficacy, 100 pharmacological properties, 102 materials and methods animals, 88 cell lines expressing GABAB receptor subunits, generation, 88 DmGB2, amino acid sequence alignment, 92f DmGB2 expression, Western blot analysis, 95f expression vectors, construction, 88 functional metabotropic GABA receptor, PeaGB1/DmGB2, 96

GABAB receptors, functional and pharmacological characterization, 89 GABAB receptor subunits, molecular properties, 91 GABA receptor subunits, amino acid identity/similarity, 95t high-affinity GABAB receptor agonists, structures, 99f immunocytochemistry, 90 PeaGB1/DmGB2, functionality and pharmacology, 97 PeaGB1/DmGB2, immunohistochemical localization, 96f PeaGB1/DmGB2, pharmacological characterization, 98f receptor-encoding cDNA clones, sources, 88 results, 90 Western blot and immunodetection, 90

R Rice planthoppers, insecticide resistance Drosophila melanogaster RDL GABA receptors, effects of A2’ mutations, 33 fipronil and ethiprole, concentration-response curves, 34f fipronil and ethiprolea, comparison of inhibitory activities, 33t ethiprole, resistance, 32 fipronil, resistance, 26 A2’N·R340Q double mutation on the GABA, influence, 31f A2’N mutation, influence, 30f fipronil, concentration-response curves, 29f GABA, concentration-response curves, 28f phenylpyrazoles, fipronil and ethiprole, structures, 27f RDL GABA receptor subunit and mutations, 27f imidacloprid, resistance, 25 organochlorines, organophosphates, resistance, 24

S Silkworm Bombyx mori, functional characterization of dopamine and

156

DA-induced luminescence, dose-response curves, 113f DA or OA receptor synthetic agonists, effects, 114 DA-stimulated luminescence, antagonist effects of DA and OA receptor ligands, 116f HEK-BmDopR2 Cells, antagonist effects, 115 HEK-293 cells, transcriptional expression of three B. mori DA Receptors, 111 HEK-mock and -BmDopR1–3 cells, RT-PCR using total RNA extracted, 112f HEK-mock and HEK-BmDopR2 cells, effects of DA or OA receptor synthetic agonists, 115f increasing concentrations of DA, effects, 113f

neuropeptide G protein-coupled receptors, 109 discussion, 117 OA receptors, DMCDM, 118 materials and methods aequorin bioluminescence Ca2+ assays, 121 aequorin vectors, construction, 120 BmDopR1–3 in HEK-293 cells, RT-PCR, 120 BmDopRs and BmETHR, expression vectors, 119 cell culture and stable transfection, 120 chemicals, 119 results biogenic amines on Ca2+-dependent luminescence, effects, 114f BmDopR1-3 by DA, Ca2+-dependent luminescence, 112 BmETH, effects, 117f BmETHR using the aequorin assay, functional analysis, 116

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