Advances in Agrochemicals: Ion Channels and G Protein-Coupled Receptors (GPCRs) As Targets for Pest Control [1] 0841232571, 9780841232570

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Advances in Agrochemicals: Ion Channels and G ProteinCoupled Receptors (GPCRs) as Targets for Pest Control Volume 1: Ion Channels and Gap Junctions

ACS SYMPOSIUM SERIES 1264

Advances in Agrochemicals: Ion Channels and G ProteinCoupled Receptors (GPCRs) as Targets for Pest Control Volume 1: Ion Channels and Gap Junctions 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 9780841232563 (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

The paper used in this publication meets the minimum requirements of American National Standard for Information Sciences—Permanence of Paper for Printed Library Materials, ANSI Z39.48n1984. Copyright © 2017 American Chemical Society Distributed in print by Oxford University Press All Rights Reserved. Reprographic copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Act is allowed for internal use only, provided that a per-chapter fee of $40.25 plus $0.75 per page is paid to the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA. Republication or reproduction for sale of pages in this book is permitted only under license from ACS. Direct these and other permission requests to ACS Copyright Office, Publications Division, 1155 16th Street, N.W., Washington, DC 20036. The citation of trade names and/or names of manufacturers in this publication is not to be construed as an endorsement or as approval by ACS of the commercial products or services referenced herein; nor should the mere reference herein to any drawing, specification, chemical process, or other data be regarded as a license or as a conveyance of any right or permission to the holder, reader, or any other person or corporation, to manufacture, reproduce, use, or sell any patented invention or copyrighted work that may in any way be related thereto. Registered names, trademarks, etc., used in this publication, even without specific indication thereof, are not to be considered unprotected by law. PRINTED IN THE UNITED STATES OF AMERICA

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.

Agrochemical Discovery - Building the Next Generation of Insect Control Agents ........................................................................................................................ 1 Thomas C. Sparks and Beth A. Lorsbach

2.

Ligand-Gated Chloride Channels and Phenolamine GPCRs Are Important Targets of Pest Control Chemicals ....................................................................... 19 Yoshihisa Ozoe

3.

Targeting Voltage-Gated Sodium Channels for Insect Control: Past, Present, and Future ................................................................................................ 37 David M. Soderlund

4.

Realizing the Potential: Improving a Microtransplantation Assay Based on Neurolemma-Injected Xenopus Oocytes .............................................................. 53 Steven B. Symington, Edwin Murenzi, Abigail C. Toltin, David Lansky, and J. Marshall Clark

5.

Recent Advances in the Functional Characterization of Honeybee Voltage-Gated Ca2+ Channels ............................................................................... 75 T. Cens, M. Rousset, J-B. Thibaud, C. Menard, C. Collet, M. Chahine, and P. Charnet

6.

Mosquito Gap Junctions: Molecular Biology, Physiology, and Potential for Insecticide Development ........................................................................................ 91 T. L. Calkins and P. M. Piermarini

7.

Glutamate Receptor-Cation Channel Complex: An Unexploited Target for Mosquito Control .................................................................................................. 111 Aaron D. Gross, Rafique Islam, and Jeffrey R. Bloomquist

8.

Metabolites of Induced Fungi: A Potential Chemical Library for Next-Generation Pesticides ................................................................................. 125 S. Furutani, M. Ihara, K. Kai, H. Hayashi, and K. Matsuda

Editors’ Biographies .................................................................................................... 133

Indexes Author Index ................................................................................................................ 137

vii

Subject Index ................................................................................................................ 139

viii

Preface On the occasion of Prof. Yoshihisa Ozoe receiving the International Award for Research in Agrochemicals from the American Chemical Society Agrochemicals Division, a symposium was organized by Aaron Gross and Joel Coats in Dr. Ozoe’s honor. There were many excellent presentations from an array of international experts in the fields of ion channels, G-protein coupled receptors (GPCRs), and gap junctions. The overall theme was movement toward a broader understanding of how endogenous biogenic amines function at various neuroreceptors and the mechanisms by which some insecticides interfere with normal function at those receptors. There were new mechanisms of action, novel chemistries, new targets and methodologies, and creative ways of attacking old targets. The opening chapter of this volume is authored by Dr. Tom Sparks who has been living and breathing insecticide mechanisms of action for many years. He is certainly one of the world’s foremost authorities on the inexorable demand for novel ways to deal with pests that threaten the security of our food, fiber, households, and public health. It is followed by the chapter from Yoshi Ozoe, the Award Winner, who provides an in-depth discussion of chloride channels and phenolamine GPCRs as important targets for insecticides currently and in the future. Dr. Ozoe’s chapter is followed by contributions by a list of the most highly regarded insecticide scientists in the world. Each of their chapters provides molecular insight into well-known targets, prospective new targets, or creative novel techniques for examining interactions between insecticides and their target sites or for evaluating compounds in search of more potent, more selective, or more environmentally friendly active ingredients. An overview of research on voltage-gated sodium channels is especially relevant given the number of historically important and current insecticides that assert their toxic actions at those sites. Other chapters address new methods for investigating action of insecticides (microtransplant techniques and gap junctions), insecticide effects on highly important species (mosquitoes and honey bees), or chemical prospecting in cultured strains of fungi. We feel that this book will provide a timely and thoughtful treatment of the current state of the art. We hope it serves as a timely update on several critically important groups of insecticide chemistry and can serve as a valuable reference volume for the readers. Each of these chapters provides many references which can inform pesticide scientists everywhere and perhaps springboard a new generation of pest management scientists, educators, and trainers.

ix

Aaron D. Gross Virginia Polytechnic Institute and State University Department of Entomology 207 Latham Hall (MC 0390) 220 Ag Quad Ln 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

Agrochemical Discovery - Building the Next Generation of Insect Control Agents Thomas C. Sparks* and Beth A. Lorsbach Dow AgroSciences, 9330 Zionsville Road, Indianapolis, Indiana 46268, United States *E-mail: [email protected]. E-mail: [email protected].

An expanding and often upwardly mobile global population requires large improvements in the quantity and quality of food production as well as freedom from the ravages of disease carrying insect vectors. This necessitates that new options and approaches for insect control be developed. Increasing pest resistance to existing insecticides, a changing regulatory landscape, and shifts in pest spectrum due to changes in climate and agronomic practices, including transgenic plants, all present challenges to developing new insect control agents. The agrochemical industry has been developing synthetic organic insecticide solutions to insect pest problems for more than 70 years. Early efforts produced just a few insecticide classes/modes of action (MoA), each with a large number of different active ingredients. More recent industry efforts have been focused on an increasingly diverse array of insecticide classes, most often coupled to new or underexploited MoAs, but with each class having only a few members. A wide range of approaches have been and continue to be employed in the discovery of these more recent commercial offerings as well as the insecticides currently under development. In spite of the decline in the number of agrochemical companies in the US, EU and Asia that are now involved in insecticide discovery, innovative solutions continue to be found. Powered by the availability of new research tools and an increase in the size of many of the remaining agrochemical companies, robust discovery platforms are being built which will provide additional novel insect control products in the future.

© 2017 American Chemical Society

Introduction The need to feed an expanding global population remains a fundamental factor in seeking improvements in crop production. Like the long standing global threat from malaria, the Zika virus outbreak in the Western Hemisphere is just the latest example of how the population can be placed in peril by insect disease vectors. Part of the equation to feeding the world and minimizing disease transmission lies in effective insect control. Although the fundamental tools and approaches to manage insect pests have been in place for some time, new options (e.g. transgenic plants, sprayable RNAi) continue to be implemented providing the growers and vector control operators with an expanding range of options (Figure 1). Although insecticides are just one option in the insect pest control toolbox (Figure 1), they remain a critical component in most integrated pest management (IPM) programs today. In a number of crops such as corn, cotton and soybeans, conventional insecticides have been replaced by transgenic plants expressing a range of toxins to control above and below ground insect pests (1). However, resistance to these insect toxins is appearing (1–3). This highlights the continuing importance and need for conventional insecticides as adjuncts to transgenic crops in many crop systems. Moreover, integrated solutions, that is traits coupled with a crop protection foliar products, can potentially provide new pest control options for growers. Alternatively, biopesticides (4, 5) present additional options for pest insect control. Although their use is on the rise, at present biopesticides represent a small component (ca 5%) of the total pesticide market (5) with conventional synthetic organic pesticides still the predominant tools in many crop production systems due to their cost, efficacy and reliability.

Figure 1. Options for insect control and delivery systems – current and potential. Options in red are innovations implemented in the past two decades or possible in the future. (see color insert) Key drivers for the development of new synthetic organic insecticides include expanding insect resistance to insecticides (2, 3) and increasingly stringent environmental, toxicological as well as regulatory requirements which limit the 2

use and/or availability of many older chemistries (6). The regulatory challenges restricting the use of older chemical insecticides also influence discovery strategies to identify new chemical pest control solutions. Those companies that can successfully navigate the evolving regulatory landscape will be rewarded by the market as new products are launched without significant delays. Likewise, changes in spectrum of insect pests in a crop as a result of changing agronomic practices, including the introduction of transgenic crops, also drive the need for new insect control options. As a prelude to understanding where agrochemical research and insecticide discovery is going it is useful to understand where the industry has come from. Thus, this review aims to provide a background on the evolution of insecticide discovery and some thoughts on directions moving forward. As will be noted later in this review, the insect nervous system has been the primary target for the majority of the insecticide classes developed in the past 70 years. In particular a range of ion channels and G-protein coupled receptors have been the targets of the neuroactive insecticides. In part, the emphasis on these types of molecular targets is due to their sensitivity to disruption by xenobiotics such as insecticides and the resulting rapid physiological cascade leading to the effective control of a wide range of pest insects.

Early Insecticide Research: Exploiting Few Modes of Action Since the introduction of the first synthetic organic insecticides more than 70 years ago, there has been a continuing evolution in the numbers, classes and modes of action of insecticidal chemistries explored and developed (Figures 2,3,4). For example, in the 1950s and into the early 1960s the primary focus was on four classes of chemistry that exploited only three different modes of action.

Figure 2. Changes in the numbers of new active ingredients introduced for the major classes of insecticides as a function of time. Chart based on data derived, in part, from the Allan Wood database (7) & Cropnosis (8). (see color insert) 3

Figure 3. Timing of the introduction and size (number of ais) of the different classes of insecticides during the past 70 years. Data derived, in part, from the Allan Wood database (7) & Agranova (9).

Commercial insecticide offerings included the organochlorine sodium channel modulators (DDT analogs), inhibitors of acetylcholinesterase (AChE) (organophosphates (OPs) & carbamates) and blockers of the gamma-aminobutyric acid (GABA) gated chloride channel (cyclodienes, BHC) (2, 7, 8). Within each class a large number of active ingredients were developed. For example, more than 150 different OP and 40 different N-methyl carbamate insecticides (Figure 3) were introduced; see also reference (2). While the first synthetic pyrethroid (allethrin) was developed in the late 1940s, the inherent photo-instability limited its use and impact. It was not until late 1970’s that the first photostable synthetic pyrethroids (e.g. permethrin, fenvalerate) were introduced for crop use. Continued commercial interest in this area of chemistry ultimately has given rise to more than 75 different active ingredients (ais) (Figure 3). Interestingly, nearly four decades after their initial introduction, new pyrethroids are still being launched today, albeit primarily for public health and vector control uses (7, 9). The late 1970s also saw the introduction of the acylureas (Figure 2) eventually totaling 14 different ais to date (Figures 2,3) (2). Shortly thereafter, the avermectins were introduced (Figure 3), which served to reaffirm the value of natural products (NPs) in the agrochemical and especially the public health arenas. Since the high water mark of the 1980s there has been an overall decline in the introduction of new insecticidal ais (10, 11). This decline is due in part to advances in intellectual property protection coupled with consolidation in the agrochemical industry and increasingly more stringent regulatory requirements (6) which have all contributed to more classes of chemistries, but with fewer members. 4

Figure 4. Timing of the introduction and size (number of ais) of the different insecticidal modes of action for synthetic organic insecticides during the past 70 years. Data derived, in part, from the Allan Wood database (7), Cropnosis (8) & Agranova (9).

Although not directly derived from the NP nicotine, the development of the neonicotinoids in the early 1990’s gave rise to what may be the most impactful class of insecticides in the past 25 years. Although comparatively few in number (presently seven different ais) compared to prior classes of insecticides, such as the OPs, carbamates, pyrethroids, and acylureas (Figure 3), the neonicotinoids captured one-quarter of the total global insecticide market in 2015 (Figures 5,6; end user sales = 24%) (9), down slightly from a high of nearly 30% in 2012 (Figure 5). The 1990s through the 2000s also gave rise to an expanding array of new classes of insecticides including the fiproles, spinosyns, METI (inhibitors of mitochondrial electron transport at complex I) acaricides, ecdysone agonists, oxadiazines, cyclic ketoenols, diamides, etc. (Figure 3), representing a range of new modes of action (MoA) (Figure 4) (2).

Current Insecticide Research: Identification of New Modes of Action In contrast to prior decades where few modes of action were utilized by the majority of insecticides in use, today more than 25 different MoAs (Figure 4) are exploited by mainstream insecticides and acaricides (2, 3), although clearly not all MoAs are available for every combination of crop-pest-location. Likewise, there are more than 30 different chemical classes of insecticides (Figures 3,4), with other new options in development (7, 11). One benefit of these new classes of chemistry is to provide new options for insecticide rotation, an important 5

approach in the management of insecticide resistance (2, 3). In addition, since the late 1990’s growers have had additional options for some pest-crop combinations in the form of transgenic plants. What has been rather interesting, but not surprising, is the virtual disappearance of the some early classes of insecticidal chemistries such as the cyclodienes, and other organochlorines, and the rather precipitous decline in global sales of the OPs and carbamates over the past two decades. These declines, in many instances, reflect regulatory agencies removing classes of chemistry with undesired toxicological and environmental profiles coupled with expanding pest resistance. Declines in sales for the above mentioned insecticide classes have been countered with rapid market penetration of new products such as the neonicotinoids, and most recently the diamide class of insecticides (Figure 5). The latter is especially interesting in that the diamides now garner a larger percentage of the global sales (10%) than either of the former mainstream insecticides, the OPs (9%) or carbamates (4%) (Figure 5). Of equal importance is the rise in sales of “other” classes of insecticides outside the OPs, carbamates, pyrethroids, and most recently the neonicotinoids, that have all been major forces in the insect control, and which clearly highlight that the market values innovation. In 1988 the OPs and carbamates together accounted for more than 60% of the total global insecticide sales and pyrethroids another 20%. Thus 80% of the market was dominated by just three classes of chemistry and two modes of action. The combined “other” classes accounted for just 11% of the sales (Figure 4). In sharp contrast, “other” classes of insecticidal compounds (including the diamides) now account for 47% of insecticide sales (Figure 4), a trend that is likely to continue.

Figure 5. A. Changes in the percentages of the insecticide market (global sales) as a function of time for selected classes of insecticides. Data from Agranova (9). B. Makeup of the Others group of insecticides based on Insecticide Resistance Action Committee (IRAC) classification (2, 3) and Agranova 2015 end user sales data (9) - millions USD. (see color insert) 6

Figure 6. Global sales [2015 end user, Agranova (9)] for the major classes of insecticides based on IRAC classification (2, 3). 50% of the global end user sales is from classes of insecticides introduced since 1990 (red & yellow). Total value in 2015 = $18.3 billion USD. Sales since 1990 are about equally divided between sap-feeding (red) and chewing (yellow) insecticides. (see color insert) As noted in Figure 3, the number of members within the more recent insecticidal chemical classes are relatively small compared to that of the carbamates (41), pyrethroids (81), acylureas (14), and especially OPs (165). Even the DDT analogs and the cyclodienes which only had 9-15 members each have not yet been matched by the most prolific insecticidal classes of the last two decades. There are currently 8 neonicotinoids, 6 METI acaricides / insecticides (Figure 3), and perhaps in the near future as many as 6 diamides. Currently flubendiamide, chlorantraniliprole and cyantraniliprole are in the market and cyclanilprole, tetraniliprole, cyhalodiamide are in the later stages of development. As noted above, in part, the smaller size of the recent new classes of insecticides is due to a number of factors including the nature of the chemistries, current chemistries being arguably more complex and more expensive, and fewer companies (US and Europe) involved in insecticide discovery than two decades ago (6, 12). Likewise, the desire for new MoAs tends to focus discovery efforts on different types of chemistries more likely to have a novel MoA. However, some of these latter factors have been mitigated in part by the expanding discovery programs of Japanese agrochemical companies (13, 14). Today, companies also tend to submit more and larger patents around an area of chemistry (6), increasing the resources required by competitors to successfully create novel compounds outside of these patents. There are also the increasing costs of discovery, registration and development of new insecticides limiting the number of chemical classes and products that a company can afford to invest in (6, 12, 15, 16). Regardless of these considerations, new chemistries, often with new MoAs, continue to be discovered and developed. Importantly, market considerations, as well as IPM and insecticide resistance management programs place a premium on new MoAs, helping to drive the search for and development of new classes of chemistries possessing these 7

new MoAs. Interestingly, in some cases very different chemistires have been discovered that are later found to address the same target site, as has been the case with the METI inhibitors fenproximate, pyridaben, fenzaquin, and tebufenpyrad (17) and, more recently, the vanilloid-type transient receptor potential (TRPV) chordotonal channel modulators pymetrozine, pyrafluquinazone and afidopyropen (18, 19). Thus, when discovered or built (see below), new classes of insecticides do not always result in new MoAs as reflected in the numbers of new molecules / classes (Figure 3) compared to the fewer numbers of new MoAs in the same time period (Figure 4). While instances of unexpected MoA repetition do limit options for rotation of MoAs for resistance management (2, 3), there can still be significant differences in spectrum, efficacy and susceptibility to metabolic resistance mechanisms, potentially minimizing the impact of the same MoA and bringing a favorable value to a new molecule.

Approaches to Insecticide Discovery – Present and Future Agrochemical discovery and the approaches that have been or could be employed have been frequently discussed (12, 15, 20–28). An examination of insecticide lead chemistries that ultimately became products since 1990 illustrates that there are a wide array of starting points/approaches for agrochemical discovery (Figure 7). In this analysis, the focus was on the new classes of chemistry (Figure 3); hence older chemical starting points (OPs, carbamates, pyrethroids, etc.) were omitted. Many of the six approaches highlighted in Figure 7 are not new and have been used in agrochemical discovery since the beginning of the insecticide revolution of the 1940s-1950s.

Figure 7. Approaches to insecticide discovery. Origins of insecticides belonging to new classes of chemistry introduced since 1990 (see Figure 3). n = 57. (see color insert) 8

1 – Competitor inspired (CI) starting points remain one of the most important sources of inspiration for new agrochemicals (Figure 7). Recent examples include the neonicotinoids (e.g. imidacloprid, thiamethoxam, clothianidin, etc.), some diamides (chlorantrianiliprole, cyclantraniliprole), oxadiazines (indoxacarb), and semicarbazones (metaflumizone). In some cases the second or third product in a class of chemistry ultimately does better, in terms of sales as critical improvements are made to the later generation ais to enhance their efficacy, spectrum, cost, regulatory or environmental profiles relative to the original product. However, CI-based discovery typically concedes points in terms of innovation, especially where a new MoA is desired, since the MoA typically mirrors the earlier molecules used as starting point(s). On occasion, however, a new MoA can arise. For example, in exploring the diamide chemistry motif, moving from an ortho-configuration to a meta-configuration for the amides resulted in a remarkable shift in MoA. The original chemistry acting at an allosteric site on the ryanodine receptor (29) evolved to a new class of insecticides (e.g. broflanilide, Figure 8) which interacts with an allosteric site on the GABA-gated chloride channel (30). Certainly, this MoA shift is the exception and not the rule. However, the above example does demonstrate that new MoAs can arise from any discovery approach. 2 – A next or second generation product derived from an existing internal product remains a validated approach to new agrochemical products and really represents a variation of the CI approach. Instead of looking externally, the research maintains an inward focus with the potential advantage of having fewer intellectual property issues than with the CI approach. The concept has been aptly demonstrated both in the past (e.g. parathion–methyl parathion; permethrin–cypermethrin–deltamethrin) and more recently (e.g. spirodiflofen–spiromesifen–spirotetramat; chloranthraniliprole–cyanthraniliprole; spinosad–spinetoram, tebufenpyrad–tolfenpyrad, abamectin–emamectin benzoate, etc.). The key, much like the CI approach, is to identify areas for possible differentiation, with efficacy and especially spectrum, being the more prominent areas for separation. 3 – A third validated approach to identify new agrochemcials is to exploit NPs (Figure 7), either directly as products (e.g. abamectin, spinosad), as semi-synthetics (e.g. emamectin benzoate, spinetoram, lepimectin, afidopyropen), or as inspiration for synthetic mimics (31, 32). NP-inspired insecticides include the aryl N-methylcarbamates (33), synthetic pyrethroids, nereistoxin analogs, chlorfenapyr, juvenoids, and more recently the butenolides (34). The use of NPs as a source of new chemistry has the advantage of being a good source of new MoAs with more than 60% of all agrochemcials MoAs having a potential NP model (31). For insecticides the value of NPs as a source of new MoAs is close to 75% (31, 32). Certainly pharmaceutical companies have leveraged NPs for inspiration as many drug products have roots in NPs (35). Moreover, there is a continuing interest in exploiting natural products for agrochemical starting points. NPs have been, or potentially could have been models for most classes of insecticides (32), with the 2015 global sales value approaching 80% of the total. However, a distinct disadvantage with NPs is that the goal of searching for a new NP 9

starting point can be more resource intensive than other approaches depending on the implementation (i.e. screening for new NPs). Also, attempting to morph a NP into an agriculturally viable molecule can require more time than other approaches (12). 4 – Compounds possessing ag-like properties or bioactive scaffolds (BAS) have also served as starting points for the discovery of more recent insecticides (Figure 7) including sulfoxaflor, pyflubumide, pyridalyl, flonicamid, and etoxazole. Also included in this broad suite of approaches are fragment, ligand, shape, and pharmacophore-based tactics (28). The focus on physical properties, Lipinski rule of 5 (36) as well as Tice ag-like properties (37) have also directed many discovery efforts to begin with a BAS or ag-like building blocks that possess the desired attributes. While there are always exceptions to these rules, in particular natural products, many of the commercial insecticides past and present fall within the ranges of ag-like properties. Moreover, as environmental factors become increasingly difficult to manage, physical properties of modern insecticides will be increasingly important. Solubility and log P in particular could be factors for ground water and soil persistence. In addition to ag-like properties, the use of novel building blocks (38) or privileged structures (39, 40) have been widely utilized in pharmaceutical drug discovery. Privileged structures are molecular frameworks that have the potential for diverse binding properties, allowing them to be developed into potent compounds for a broad range of biological targets. Moreover, privileged structures can have ag-like properties at the start which accelerates optimization. Recent efforts to focus on developing chemical reactions and transformations that deliver polar motifs have been showcased by AstraZeneca (41). The need for smaller, polar starting scaffolds, perhaps with chirality will open new chemical space that has yet to be exploited in agrochemical discovery in general and certainly for insecticide research. 5 – Broad screening of internal chemistries coupled with data-mining (Figure 7) has also been an effective means to identify new leads that in turn have been developed as products. Bifenazate, spirodiclofen, tebufenozide, fipronil, and tebufenpyrad are all examples of recent insecticides with origins in other therapeutic areas (e.g. herbicides) that were identified by screening all chemistries broadly. Likewise, the initial lead chemistry for the isoxazolines (e.g. fluxametamide, Figure 8) appears to have also come from the broad screening of synthetic intermediates. 6 – Among the approaches employed for insecticide discovery (Figure 7), screening 3rd party compounds for insecticidal activity would appear to be the least favored in the last two decades. However, these 3rd party inputs have had a very large impact on the agrochemical industry, especially for insecticides. It was a 3rd party input that led to the discovery of nithiazine (42) which in turn led to the discovery of imidacloprid (43) and the beginnings of the neonicotinoid class of insecticides (44), currently the largest class of insecticides in terms of global sales (24%, Figures 5,6). Likewise, a 3rd party herbicidal diamide lead chemistry gave rise to flubendiamide and the diamide class of insecticides (45) which now accounts for 10% of the global market (Figures 5,6). The value of 3rd party compound inputs can be further enhanced through careful hypothesis testing 10

and the use of chemoinformatic tools to narrow and focus the inputs towards more ag-like or lead-like chemistries (27, 28). The above approaches to insecticide discovery can also be enhanced or augmented through the use of in vitro screening and/or the application of quantitative structure activity relationships (QSAR). In vitro screening has been examined as a tool for insecticide discovery for quite some time, and remains of interest (46). Genomics also presents opportunities to identify new targets of interest. However, the translation of in vitro target-site inhibition to the whole organism bioactivity, to commercial levels of field efficacy, has proved rather elusive. Some success was observed in very early insecticide discovery efforts where in vitro screening was part of discovery programs for aryl-N-methyl (propoxur (33),) and later N-methyl oxime (aldicarb (47),) carbamates. Likewise, QSAR has been widely used as a tool to understand insecticide action and efficacy after the fact, but somewhat surprisingly, only rarely has QSAR been the driving force in the insecticide discovery process. Two such examples where QSAR has played a pivotal role in insecticide discovery include the discovery of the pyrethroid bifenthrin (48) and the next generation spinosyn insecticide, spinetoram (49).

Summary and Conclusion As noted above, in the early days of synthetic organic insecticides, there were only a few modes of action that were widely exploited. Current trends and data suggest that a new MoA is and will continue to be an important attribute for a new insecticide. Since 1990 a number of new classes of insecticides have been developed (Figure 3) that possess new modes of action (Figure 4), and these new classes presently account for 50% of the global agrochemical sales (Figure 6). Pesticide discovery is now a global enterprise spread increasingly across companies in the US, EU and Asia (esp. Japan and China). As outlined above, there also continues to be an array of new insecticides and MoAs being brought to the market (Figures 3,4). However, notwithstanding these advances in insecticides and acaricides, there remains a pressing need for new insect and mite control options, since (as also noted above) some crop-pest-location complexes may have very few options. There are several new sap-feeding insecticides in development, some exploiting known MoAs (e.g. afidopyropen (19);), while the MoA of others (fluhexafon, benzpyrimoxan, Figure 8) is currently unknown. Their commercialization could be especially important to fill a market void should any reduction in the use of neonicotinoids, which presently hold a large share of the insecticide market, occur due to resistance, regulatory aspects or public perception. During the past two decades, the agrochemical industry has incorporated an array of tools including combinatorial chemistry (50, 51), high throughput screening (52), and structure-based design (15), all focused on enhancing agrochemical discovery. Random screening of compounds has generally given way to more targeted efforts to bring in compounds that fit some a priori hypothesis (15, 28) or have been pre-filtered for agrochemical lead-like properties 11

or computational likeness for molecules targeted for pest group (52). The ‘omics revolution presents opportunities to better understand insecticide MoA or envision new potential target sites for new insecticides. However, as with any new technology, hope and hype runs high, but thus far, none of these past or present techniques have proven a panacea for insecticide discovery. Nevertheless, these new technologies have made the discovery process more efficient and have proven useful in the exploration and exploitation of new insecticidal leads. Likewise, MoA determination now has more tools available to potentially speed the process (53). These tools can be further enhanced by collaboration with academic research groups that can bring greater focus or expertise to MoA determination to allow the more problematic MoAs to be successfully resolved. This concept has been aptly demonstrated by the recent elucidation of the MoA of pymetrozine through a collaboration between an agrochemical company and a university (18).

Figure 8. Structures of recent insecticides. Regardless of the approach taken, it is self-evident that the best place to look for biological activity is in other biologically active molecules. Hence the continued interest in NPs, and the long term overriding interest in CI as a means to discover new insecticides. Likewise, broad screening of chemistries from other therapeutic areas has frequently identified insecticidal analogs that have served as leads for future products. The fiproles, cyclic ketoenols, formamidines, and diamides all had their origins in herbicidal chemistries. Thus, it is reasonable to assume that other sources such as pharmaceuticals can also give rise to new insecticides. In many respects the physical properties of pharmaceuticals and agrochemicals are not dissimilar (54, 55), and it has also been suggested that agrochemcials may, in turn, provide starting points for new drugs (54). 12

The fundamental approaches (as outlined above) to find that initial discovery have remained constant for at least the last three decades, but the terminology and tools available have evolved with time, and have contributed to delivery of new insecticidal products. Computer aided molecular design (CAMD), QSAR and chemoinformatics can aid in that task providing the tools to more easily uncover the next new insecticide lead. However, the essential catalyst in insecticide discovery process has been and will remain the scientist(s) asking new questions that can drive innovation. Importantly, new insecticides / agrochemicals are not so much “discovered” as they are “built”. What does this mean? Typically starting with a screen hit, idea, NP, inspiration from a competitor molecule, model, etc., hypothses are formed to address a liability or limitation of the starting point, most often in the early phases, efficacy. This involves several to many iterations where a succession of analogs are evaluated until the ai with the best combination of efficacy, toxicology, cost, physical properties and environmental parameters emerges. Thus, most new agrochemicals are built substituent by substituent, sometimes atom by atom. At times this optimal analog is immediately obvious, other times it is only recognized hundreds of analogs after the fact when the summation of attribute data becomes available. It is in these instances where CAMD, QSAR and chemoinformatic tools can expedite the process by providing an effective means to summarize / visualize the data enabling the optimal analog and potential future insecticide to be identified.

Acknowledgments We thank Mr. Greg Hanger (DAS), Drs. Ronda Hamm (DAS), Jim Hunter (DAS), Frank Wessels (DAS), Debra Camper (DAS), John Casida (University of California, Berkeley), and Rob Bryant (Agranova) for their very helpful discussions and comments.

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Chapter 2

Ligand-Gated Chloride Channels and Phenolamine GPCRs Are Important Targets of Pest Control Chemicals Yoshihisa Ozoe* Faculty of Life and Environmental Science, Shimane University, Matsue, Shimane 690-8504, Japan *E-mail: [email protected].

Ion channels and G protein-coupled receptors (GPCRs) are important targets of pest control chemicals. Research on GABA-gated chloride channels (GABACls) identified the firstand second-generation of noncompetitive antagonist (NCA) insecticides in the 1980s. The resistance of insect pests to these insecticides has recently become a serious concern. However, the recent development of the third-generation of NCA insecticides, which have distinct mode(s) of action, overcomes this problem and expands the potential of GABACls as multi-site targets. Despite decades of efforts to discover pest control chemicals targeting octopamine GPCRs, no practical agent has been developed since chlordimeform and amitraz. However, given that different types of phenolamine GPCRs were recently identified in various invertebrates, the detailed analysis of these receptors, in particular, β-adrenergic-like octopamine receptors with high sensitivity to agonists and a widespread distribution in the body, may provide new opportunities to create second-generation octopamine receptor agonist insecticides.

Ion channels play vital roles in neurotransmission in the nervous system, and invertebrate ion channels are important targets for pest control chemicals (1). These channels include voltage-dependent sodium channels, nicotinic

© 2017 American Chemical Society

acetylcholine-gated cation channels, γ-aminobutyric acid (GABA)-gated chloride channels (GABACls), L-glutamic acid (glutamate)-gated chloride channels (GluCls), and calcium-activated calcium channels as the targets of existing insecticides (2). Transient receptor potential (TRP) cation channels have recently been added to this group (3). G protein-coupled receptors (GPCRs) constitute the largest family of membrane receptors. More than 800 and 200 GPCRs are encoded in the human and Drosophila genomes, respectively, and they are involved in a variety of physiological processes (4). It has been argued that insect GPCRs are attractive targets for insecticides (5, 6), although octopamine receptors are the only GPCRs currently targeted by existing acaricides. This article reviews our long-term research on ligand-gated chloride channels and phenolamine GPCRs and shows their importance as the targets of pest control chemicals.

GABA- and Glutamate-Gated Chloride Channels Early-Stage GABA Receptor Research Our research on GABA receptors started in 1976 when we encountered ethylbicyclophosphate (1) (Figure 1), which had been identified as a toxic combustion product from a fire-retarded polyurethane foam. This type of unique phosphate is relatively stable and does not react with acetylcholinesterase (7), which is the target of organophosphorus insecticides. We first synthesized a number of analogs to obtain clues for its mode of action. This class of compounds also exhibited, albeit low, insecticidal activity by injection (8). We performed quantitative structure-activity relationship (QSAR) analyses using the Hansch-Fujita method (9). The difference in the coefficient of the steric parameter Esc of the bridgehead substituent between regression equations for toxicity in mice and houseflies (Musca domestica) allowed us to speculate that the bicyclophosphate binding sites of mice and houseflies might have structural differences, which would be the basis of the development of safer insecticides (7).

Figure 1. Chemical structures of noncompetitive GABA receptor antagonists. 20

In 1977, we reported the effects of bicyclophosphates on the miniature inhibitory junction potentials observed in the longitudinal muscle of the earthworm (Metaphire communissima) (10). When the isopropyl analog (IPPO) at 10 μg/ml was applied onto a muscle preparation, the spontaneous discharges declined with time and were finally eliminated. This muscle is innervated by neurons containing the inhibitory neurotransmitter GABA. This observation implied that the antagonism of GABA receptors underlies the toxic action of bicyclophosphates. While synthesizing bicyclophosphates, we found that an analog (TBPO) with a tertiary butyl group is highly toxic to mice, with an LD50 of 0.053 mg/kg, which was determined by intraperitoneal injection (9). We aimed to identify the binding protein(s) using radiolabeled TBPO. Although our labeled compounds were unsatisfactory because of low specific activity (7), the thiono analog (TBPS) (2) (Figure 1) of TBPO was successfully labeled with 35S by Squires and coworkers (11) and found to be an excellent GABA receptor probe (12), along with [3H]4′-ethynyl-4-n-propylbicycloorthobenzoate (EBOB) (3) (Figure 1) (13). In 1982, Ghiasuddin and Matsumura reported that cyclodienes and γ-BHC act as GABA receptor antagonists (14). The structural similarity of heptachlor epoxide and γ-BHC to the GABA receptor antagonist picrotoxinin (Figure 1) is one of the lines of evidence for this claim. However, these compounds look different from cage compounds, such as TBPS and EBOB. To probe the binding site, we synthesized hybrid compounds (4) with a variable number of chlorine atoms in the ring and various bowsprit substituents (R) (Figure 1) (15–17). We performed 3D-QSAR analyses of these compounds using comparative molecular field analysis (CoMFA), which allowed us to predict the differences in the structures of noncompetitive antagonist (NCA) binding sites between houseflies and rats (18). Bicyclophosphates with the appropriate alkyl groups at both the bridgehead and the endocyclic methylene are an example, demonstrating that the results from our CoMFA analysis can be utilized for the design of NCAs with selectivity for insect GABA receptors (19). Bicyclophosphates with a substituent only at the bridgehead, such as TBPO and TBPS, are highly toxic to mammals but are not highly insecticidal. However, PS-14 (5) (Figure 1), a 3,4-dialkylbicyclophosphorothionate, potently inhibited GABA responses in dissociated neurons from the American cockroach (Periplaneta americana), whereas it was an approximately 200-fold less potent inhibitor of the GABA responses in rat cerebral cortex neurons (20) (Figure 2). The introduction of an isopropyl group to the position next to the oxygen atom conferred insect receptor selectivity to bicyclophosphates. This is consistent with the CoMFA prediction that electropositive substituents near the heteroatoms within NCA structures are advantageous to high insecticidal activity (18).

21

Figure 2. Dose-response curves of the PS-14 inhibition of GABA- and glutamate-induced currents in P. americana neurons and GABA-induced currents in rat neurons. The data are presented as the mean ± SD of three experiments. (Reproduced with permission from reference (20). Copyright 2013 Elsevier.) A variety of GABA receptor antagonists have been identified to date. Picrotoxinin was the first antagonist discovered in 1956 (21). Our group has identified several classes of naturally occurring GABA receptor antagonists, including picrodendrins (22), seco-prezizaanes (23), and spiroquinazolines (24). In addition to the naturally occurring compounds, the compounds that our lab synthesized added to the NCAs (25–28). Phenylpyrazoles, which have no structural relationship to the NCAs synthesized at the time, were commercialized as second-generation GABA receptor NCA insecticides (29). Molecular Pharmacology of GABA Receptors When we began researching GABA receptors, the GABA receptor structure was unknown. GABA receptor research was facilitated by the first structural elucidation of bovine and Drosophila GABA receptor subunits (30, 31). At the present, we have much knowledge concerning GABA receptors. Mammalian GABA receptors are heteromeric chloride channels with five subunits selected from among 19 members, and insect GABA receptors are homomeric chloride channels composed of five subunits termed Rdl (32, 33). Therefore, GABA receptors are also designated GABACls in this review when mentioned in comparison to GluCls. Chloride channels are opened by GABA binding to the extracellular subunit interfaces to cause intracellular hyperpolarization due to chloride influx. Picrotoxinin blocks the channel by binding to a site within the channel. In mammalian GABA receptors, the anti-anxiety drugs benzodiazepines bind to the extracellular subunit interface to enhance GABA responses. The sedative hypnotics barbiturates and the anesthetics etomidate and propofol allosterically activate or potentiate currents in GABA receptors by binding to the transmembrane subunit interfaces. Thus, GABA receptors have multiple sites for ligands. 22

We identified the binding site of the NCA EBOB by examining the binding of [3H]EBOB to homo-pentameric human β3 GABA receptor mutants expressed in S2 cells (34). This receptor was used as a pharmacological surrogate for insect GABA receptors (35). We substituted pore-lining amino acids at the -1′, 2′ and 6′ positions of each subunit on the cytoplasmic side within the channel (the conserved TM2 arginine was index numbered 0). The substitution of the 6′ threonine with valine abolished [3H]EBOB binding, and the substitution of the 2′ alanine with serine caused a decrease in the affinity of EBOB for the receptor. These findings suggest that the hydrogen bonds between the EBOB oxygen atoms and the 6′ threonine hydroxyl hydrogen atoms may exert profound effects on EBOB binding (Figure 3) (34, 36).

Figure 3. The docking of EBOB into the channel pore of the human β3 GABA receptor modeled using PDB entry 1OED as a template. (Reproduced with permission from reference (36). Copyright 2007 American Chemical Society.)

As described above, PS-14 is selective for insect GABA receptors. This selectivity could be primarily explained by the difference in the 2′ amino acids (37). The NCA binding site is located around the 2′ and 6′ positions within the channel. PS-14 has two hydrophobic alkyl groups that fit into the space created by five 2′ alanines in the P. americana GABA receptor channel (Figure 4A). The P. americana inhibitory glutamate receptor (GluCl) and rat α1β2γ2 GABA receptors have 2′ amino acids with polar (Ser) or bulkier (Val) side chains, which most likely result in unstable PS-14 binding (Figure 4B). 23

Figure 4. Dependence of the potency of PS-14 on the 2′ amino acid. (A) The docking of PS-14 into a human β3 GABA receptor homology model. Five TM2 α-helices are shown by blue lines. (Reproduced with permission from reference (37). Copyright 2010 John Wiley and Sons.) (B) The 2′ amino acid arrangements of the P. americana GABACl, P. americana GluCl, and rat GABACl.

The 2′ and 6′ amino acids play critical roles in the interaction of NCAs with GABA receptors. Substitution of the 2′ amino acid is responsible for the resistance of insect pests to NCA insecticides (Figure 5). The A2′S mutation causes a marked reduction in the sensitivity of the receptor to dieldrin but not to phenylpyrazoles (36, 38). The wild-type common cutworm (Spodoptera litura) was shown to have a 2′ serine, which is the type of residue that confers dieldrin-resistance (39). The A2′N mutation results in a pronounced reduction in the sensitivity of the receptor to fipronil (36, 40, 41). Moreover, certain arthropod species have multiple copies of the Rdl genes that encode different 2′ amino acids. These amino acid changes cause insensitivity to conventional NCA insecticides. This most likely depends on the structural moiety of NCAs that interacts with the 2′ amino acids, which might cause the difference in the sensitivity of the receptor even to the same class of insecticides, fipronil and ethiprole (42). 24

Figure 5. The amino acid sequences of the TM2 regions of fruit fly, small brown planthopper, common cutworm, and two-spotted spider mite Rdl subunits.

GABA Receptors versus Inhibitory Glutamate Receptors Insects have homologous ligand-gated chloride channels, GABACls and GluCls. Both GABACls and GluCls generally mediate inhibitory neurotransmission, although this depends on the intra- and extracellular chloride concentrations, which are regulated by cation chloride cotransporters. It has yet to be determined why two homologous chloride channels are needed for insect physiology and which channel is toxicologically important. To address these issues, we examined the localization and expression level of the transcripts and proteins of the GABACl Rdl and the GluCl subunits. qPCR analyses showed that both Rdl and GluCl transcripts were predominantly expressed in the adult head compared with other developmental stages and other body parts (43). The Rdl and GluCl transcripts were equally expressed in most body parts, although the GluCl transcript was expressed at higher levels than the Rdl transcript in the adult prothoracic leg. We then immunocytochemically examined the localization of the subunit proteins using specific antibodies (43). Frontal sections through the adult housefly head revealed specific Rdl and GluCl staining in the optic lobe neuropils and brain regions. The GluCls were specifically found in the pigment cells, retina basement membrane, and lamina, whereas Rdl staining was intense in the medulla, lobula, and lobula plate. Rdl staining was observed in the mushroom body, antennal lobe, and central complex. In the thorax, GluCl staining was located in the monopolar cell bodies of motor neurons. In contrast, Rdl staining was rather widely distributed in the thoracic ganglion. GluCl staining, but not GABACl staining, was also observed along the femoral muscles of the prothoracic leg. The differential distribution of GABACls and GluCls was thus revealed. We expressed housefly GABACls and GluCls in Xenopus oocytes and compared their functions and the potencies of three representative NCAs against the two channels using two-electrode voltage clamp (TEVC) electrophysiology (44). Although there is no obvious functional difference between these channels, 25

the GABACls was found to be more sensitive by two to four orders of magnitude to picrotoxinin, γ-BHC, and fipronil than GluCls. All the NCAs we have tested so far exhibited selectivity for GABACls. It has yet to be clarified whether GluCls could also be important targets of NCAs. We recently examined the actions of the macrolide ivermectin on GluCls and GABACls expressed in oocytes in detail (45). Ivermectin elicited three responses in both channels: alone, it activated chloride current into the cell; and it potentiated or antagonized agonist-induced chloride currents through the channels depending on the agonist concentration. In all these responses, particularly, activation, ivermectin’s effects were more potent for GluCls compared to GABACls. The mechanism of this selectivity remains to be investigated.

New-Generation GABA Receptor NCAs Fluralaner (BravectoTM) (Figure 6) is a recently developed isoxazoline ectoparasiticide. We investigated the action of this compound on housefly GABACls and GluCls expressed in Xenopus oocytes using a TEVC technique. In 2010, we reported that fluralaner inhibits both GABA- and glutamate-induced currents in these channels (46). However, the GABACls were approximately 20-fold more sensitive to fluralaner than GluCls. We generated GABACl mutants with 2′ amino acid substitution to examine whether fluralaner shares a site of action with fipronil. The A2′N GABACl mutant exhibited low sensitivity to fipronil, whereas it was sensitive to fluralaner (38, 41). This finding indicates that fluralaner is a novel NCA insecticide that binds to a different site than fipronil. Fluxametamide (Figure 6) has recently been added to the isoxazoline class of insect pest control chemicals.

Figure 6. Chemical structures of fluralaner and fluxametamide.

We also examined the action of the benzamide insecticide BPB1 (Figure 7) on housefly GABACls and GluCls (47). BPB1 selectively inhibited GABA-induced currents in GABACls. Interestingly, conventional NCAs enhanced the specific binding of [3H]BPB1 to housefly head membranes, whereas BPB1 and GABA inhibited the binding. [3H]BPB1 equally bound to the head membranes of the wild-type and A2′S mutant houseflies. These findings indicate that the benzamides are a novel class of NCA insecticides that are different from conventional ones. Broflanilide (Figure 7) is a novel insecticide that belongs to this class of chemistry (48). The binding site was predicted to be located in the transmembrane subunit interface of GABACls (49). 26

Figure 7. Chemical structures of BPB1 and broflanilide.

Orthosteric Ligands of GABA Receptors Bicuculline and gabazine (SR95531) (Figure 8) are representative mammalian competitive GABA receptor antagonists. In contrast, the competitive antagonism of insect GABA receptors remains poorly understood. Bicuculline and gabazine are inactive or only weakly active in insect GABA receptors. We synthesized some gabazine analogs and tested them against insect GABA receptors. We found that the antagonism of insect GABA receptors was enhanced by the structural modification of gabazine (50). We examined the action of 5-(4-piperidyl)-3-isoxazolols on GABA receptors from three insect species by (51). 4-(3-Biphenylyl)-5-(4-piperidyl)-3-isoxazolol (3B-4-PIOL) (Figure 8) exhibited competitive antagonism for the GABA receptors of the common cutworm and the housefly, with low micromolar IC50s. Interestingly, this compound acted as a partial agonist against the GABA receptors of the small brown planthopper (Laodelphax striatellus). 3B-4-PIOL docked to the orthosteric sites of the GABA receptor homology models of three insect species (51). The orientations of 3B-4-PIOL varied between the different GABA receptors, but the interactions of 3B-4-PIOL with arginine and glutamic acid in the orthosteric site were a common feature of the docking (Figure 9). For the common cutworm and housefly GABA receptors, for which 3B-4-PIOL exhibited antagonism, a cation-π interaction was observed between the biphenylyl group and an arginine. In the small brown planthopper GABA receptor, for which 3B-4-PIOL exhibited agonism, the corresponding intermolecular electrostatic interaction was absent because of an intramolecular cation-π interaction within 3B-4-PIOL. This difference could explain whether this compound elicits antagonism or agonism.

Figure 8. Chemical structures of GABA and competitive antagonists. 27

Figure 9. Predicted interaction between 3B-4-PIOL and the amino acid residues in the orthosteric site of insect GABACls. Further structural modification is needed for the development of potent and selective competitive antagonists or agonists. The competitive antagonists may be lead compounds for insecticides as their action results in an allosteric blockade of the channel.

Phenolamine GPCRs Octopamine/Tyramine GPCR Subtypes The phenolamine octopamine, which is synthesized from tyramine, has been extensively studied over the past four decades from both physiological and pharmacological perspectives (52). Octopamine is a multifunctional signaling molecule that is involved in a variety of physiological and behavioral processes, and tyramine is also now considered a signaling molecule. The actions of octopamine and tyramine are mediated by the activation of specific GPCRs (52). Octopamine receptors have long been known to be the targets of formamidine acaricides and insecticides (53). N-Demethylated chlordimeform (DMCDM) was shown to be a potent agonist of octopamine receptors in assays using tissue or membrane preparations. Recent progress in cloning and molecular biology technologies has made it possible to characterize individual receptors. We identified four phenolamine GPCRs in the silkworm (Bombyx mori) as a lepidopteran model insect: (i) the first tyramine receptor, BmTAR1, which is phylogenetically close to the human α2-adrenergic receptor (54); (ii) the second tyramine receptor, BmTAR2, which is distant from human adrenergic receptors (55); (iii) the first octopamine receptor, BmOAR1, which is phylogenetically close to the human α1-adrenergic receptor (56); and (iv) the second octopamine receptor, BmOAR2, which is similar to the β-adrenergic receptor (57). We examined the localization of phenolamine receptor transcripts in the silkmoth and silkworm bodies (58). The transcript of the β-adrenergic-like octopamine receptor BmOAR2 was predominantly expressed in the brain, nerve cord, antennae, pheromone gland, oviduct, and flight muscles. Among these body parts, the BmOAR2 transcript was most abundant in the flight muscles, followed 28

by the oviduct, brain, and nerve cord. The BmOAR2 transcript was detected in the silkworm brain, nerve cord, Malpighian tubule, silk gland, and midgut. We performed a functional characterization of four phenolamine GPCRs by stably transfecting their cDNAs into HEK-293 cells. In HEK-293 cells expressing BmTAR1, tyramine attenuated the forskolin-stimulated cAMP levels with a potency greater by one or two orders of magnitude than octopamine (54). Although BmTAR1, initially named B96Bom, was considered to be an octopamine receptor, we demonstrated that this receptor is a GPCR for tyramine and that tyramine is not only a precursor for the synthesis of octopamine but a genuine signaling molecule. Yohimbine is a competitive antagonist as it caused a rightward shift of the tyramine dose-response curve. The octopamine receptor agonist DMCDM had no effect on BmTAR1 (59). Site-directed mutagenesis experiments demonstrated that the substitution of amino acids in the third and fifth transmembrane domains (TM3 and TM5), which are conserved in adrenergic receptors, eliminated or reduced the agonist activity of tyramine. An aspartic acid in TM3 and two serines in TM5 were suggested to be involved in the interaction with tyramine (60). The second tyramine receptor, BmTAR2, is most likely coupled to a Gq protein to mobilize calcium ions from the endoplasmic reticulum in response to tyramine (55). Octopamine and dopamine were very weak agonists for this receptor, and yohimbine was a potent antagonist. The effects of formamidines on BmTAR2 have yet to be investigated. Octopamine elicited calcium and cAMP signals by acting on the α-adrenergic-like octopamine receptor BmOAR1 expressed in HEK-293 cells (56). Calcium ions were mobilized by lower concentrations of octopamine relative to cAMP level elevation. Octopamine possibly interacts with conserved amino acids in TM3, 5, and 6. Interestingly, the substitution of a TM6 tyrosine with phenylalanine eliminated cAMP signaling but not calcium signaling, which indicates the importance of the hydroxyl group of tyrosine for cAMP signaling only (61, 62). We hypothesized that BmOAR1 activation occurs in two steps. When at low concentrations, octopamine most likely converts the inactive state to a semi-active state by interacting with an aspartic acid (TM3) and a serine (TM5), which results in the elevation of intracellular calcium levels. High concentrations of octopamine might shift the semi-active state to a fully active state by additionally interacting with a tyrosine in TM6, which triggers the elevation of the intracellular cAMP levels. We tested representative insecticidal agonists against BmOAR1 (63). In our analysis of cAMP signaling, DMCDM and a phenyliminoimidazolidine agonist (NC-5) were more potent agonists than octopamine. However, DMCDM was less potent than octopamine for activating calcium signaling. Clonidine was the most potent synthetic agonist, followed by naphazoline and tolazoline. We also examined the effects of antagonists on BmOAR1. Yohimbine and chlorpromazine inhibited both cAMP and calcium signaling. The β-adrenergic-like octopamine receptor BmOAR2 upregulated intracellular cAMP levels in response to octopamine (57). The R enantiomer, which is endogenous octopamine, exhibited higher potency than the S enantiomer, with an EC50 value of approximately 1 nM. DMCDM exhibited higher potency 29

than octopamine, with an EC50 value of approximately 100 pM. Naphazoline was a potent synthetic agonist, and the BmOAR1 agonist clonidine was less potent. Chlorpromazine was an antagonist of BmOAR2. Site-directed mutagenesis and docking simulations indicated major roles for three conserved amino acids in octopamine binding (64). An aspartic acid in TM3 may interact with both the amino and side-chain hydroxyl groups of octopamine. A serine in TM5 and a tyrosine in TM6 may interact with the phenolic hydroxyl group of octopamine.

Amitraz Amitraz is the only octopamine receptor agonist that is currently used for pest control. However, the resistance of cattle ticks to amitraz has become a serious problem. Amitraz is thought to be a proinsecticide that is converted to the active metabolite N2-(2,4-dimethylphenyl)-N1-methyformamidine (DPMF). We recently examined the effects of amitraz and DPMF on the calcium and cAMP signaling of BmOAR1 and the cAMP signaling of BmOAR2 (58) (Table 1). Although DPMF was actually the most potent agonist for all signal transductions, amitraz also exhibited agonist activity. Among these effects, the cAMP signaling of BmOAR2 was most strongly affected by DPMF, with a low picomolar EC50 value. Given the wide distribution of BmOAR2 and the high potency of formamidines for BmOAR2, β-adrenergic-like octopamine receptors are attractive targets for pest control chemicals. The use of heterologous expression systems for individual subtypes may aid in discovery of novel compounds that target phenolamine GPCRs. Polymorphic analysis of the tick β-adrenergic-like octopamine receptor may help determine whether the resistance is due to target site insensitivity.

Table 1. EC50 Values of Octopamine, Amitraz, and DPMF in Producing Second Messengers in HEK-293 Cells Expressing BmOAR1 and BmOAR2 EC50 (nM) Compound

BmOAR1

BmOAR2

Ca2+

cAMP

cAMP

Octopamine

23.4 (19.7-28.8)a

3030 (1970-4760)

5.23 (3.87-6.93)

Amitraz

9.62 (8.11-11.39)

708 (518-1042)

2.04 (1.29-3.47)

DPMF

1.17 (0.83-1.60)

181 (127-271)

0.0796 (0.0600-0.1084)

a 95% confidence interval in parenthesis. SOURCE: Reproduced with permission from reference (58). Copyright 2017 John Wiley and Sons.

30

Conclusions and Prospects Recent discoveries related to novel antagonist chemistry have expanded the potential of GABA receptors as important targets of pest control chemicals. Given that there are multiple sites for ligands in GABA receptors, discovery of novel chemistry is expected. Several types of phenolamine GPCRs have been identified. Although no pest control agent targeting GPCRs has been marketed after chlordimeform and amitraz, recent advances in understanding the molecular structures and functions of GPCRs will provide new opportunities for the discovery of novel pest control chemicals.

Acknowledgments I am deeply grateful to the late Professors M. Eto and F. Matsumura for their long-standing mentoring and encouragement. I also thank Drs. K. Tanaka, M. Akamatsu, and K. Matsuda for helpful discussion and collaboration. I acknowledge F. Ozoe for continuing to contribute to my research. I also acknowledge the past and present members of my laboratory for their excellent work.

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60. Ohta, H.; Utsumi, T.; Ozoe, Y. Amino acid residues involved in interaction with tyramine in the Bombyx mori tyramine receptor. Insect Mol. Biol. 2004, 13, 531–538. 61. Hamasaki, T.; Ozoe, F.; Ohta, H.; Enomoto, K.; Kataoka, H.; Sawa, Y.; Hirota, A.; Ozoe, Y. Identification of critical structural determinants responsible for octopamine binding to the α-adrenergic-like Bombyx mori octopamine receptor. Biochemistry 2007, 46, 5896–5903. 62. Hamasaki, T.; Ozoe, F.; Ozoe, Y. Single amino acid of an octopamine receptor as a molecular switch for distinct G protein couplings. Biochem. Biophys. Res. Commun. 2008, 371, 610–614. 63. 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. 64. Chen, X.; Ohta, H.; Sasaki, K.; Ozoe, F.; Ozoe, Y. Amino acid residues involved in the interaction with the intrinsic agonist (R)-octopamine in the β-adrenergic-like octopamine receptor from the silkworm Bombyx mori. J. Pestic. Sci. 2011, 36, 473–480.

36

Chapter 3

Targeting Voltage-Gated Sodium Channels for Insect Control: Past, Present, and Future David M. Soderlund* Department of Entomology, Cornell University, Geneva, New York 14456, United States *E-mail: [email protected].

Voltage-gated sodium channels play a fundamental role in neuronal signaling. The crucial importance of sodium channels is evident in the great structural and pharmacological variety of naturally-occurring neurotoxins produced by plants, animals and microorganisms that disrupt channel function, thereby contributing to the chemical warfare of predation and defense. The importance of sodium channels as insecticide targets was first established by the discovery of the natural insecticide pyrethrum more than two centuries ago. Since then, the empirical search for new insecticidal agents has "rediscovered" the sodium channel as a target many times, exploiting not only the receptor site for pyrethrum constituents and their synthetic analogs, the pyrethroids, but also other receptor sites on the sodium channel protein. Here I reflect on the history of sodium channel exploitation for insect control and the current status of sodium channel-directed insecticides, and I consider the durability of sodium channels as targets for the continued development of insect control agents.

Voltage-Gated Sodium Channels Voltage-gated sodium channels (VGSCs) are both ubiquitous and essential (1). They are located in the cell membranes of neurons and vertebrate skeletal and cardiac muscle, where they play critical roles in electrical signaling. Sodium channels open transiently in response to changes in the electrical potential across © 2017 American Chemical Society

the cell membrane, allowing sodium ions to flow into the cell. The resulting transient depolarization underlies the nerve or muscle action potential. The essential functions of VGSCs are intrinsic to a single, large, conformationally flexible transmembrane protein (the α subunit) that forms a sodium-selective ion pore (2). The structures of VGSC α subunits are highly conserved across the animal kingdom, especially in the 24 transmembrane helices and four pore-forming loops. In this regard, VGSCs differ from ligand-gated ion channels (i.e., neurotransmitter receptors). Ligand-gated ion channels are heteromultimers composed of various combinations of subunit proteins that are less conserved, at the amino acid sequence level, than VGSC α subunits. Insects and vertebrates have taken different routes to achieving functional diversity of VGSCs in different cells and tissues and at different developmental stages. In insects VGSCs are encoded by a single gene (para in Drosophila melanogaster and its orthologs in other species) that undergoes alternative splicing and RNA editing at multiple sites to yield a large theoretical number of splice and editing variants, not all of which are found in surveys of mRNA diversity (3). The functional characterization of splice variants isolated from a single species confirmed that alternative splicing yields VGSCs with different functional properties. Additional functional diversity may be conferred by coexpression in the cell membrane of the VGSC α subunit with one of a family of small auxiliary subunits. By contrast, VGSC α subunits in mammals comprise a family of nine isoforms (designated Nav1.1 – Nav1.9) that exhibit unique patterns of developmental and anatomical expression and varied functional and pharmacological properties (2). Mammalian VGSC α subunits are highly conserved among themselves (>90% amino acid identity). Additional heterogeneity among sodium channels expressed in vivo results from their coassembly in the nerve membrane with one or two small auxiliary β subunit proteins, which are structurally unrelated to the auxiliary subunits of insect VGSCs.

Coevolutionary Exploitation of Sodium Channels Sodium channels are the site of action of a large structural variety of naturally-occurring neurotoxins (4, 5). This coevolutionary convergence of strategies for chemical predation and defense on a single protein underscores both the fundamental significance of VGSCs in the normal function of animal nervous systems and the disruptive potential of agents that modify normal VGSC function. The sites of action of these neurotoxins, identified in functional and radioligand binding assays and by site-directed mutagenesis, identify seven distinct binding domains for naturally-occurring neurotoxins on the VGSC α-subunit protein (Table 1). These binding sites are operationally defined by competitive interactions among toxins considered to act at the same domain and allosteric interactions between toxins acting at different domains. However, different residues in the sodium channel protein may be critical for the binding of different ligands that compete for occupancy of same binding domain. Moreover, 38

additional neurotoxins not shown in Table 1 modify VGSC function by action at binding domains that are distinct from Sites 1-7 but otherwise incompletely characterized.

Table 1. Neurotoxin Binding Sites on Voltage-Gated Sodium Channelsa

a

Site

Active Neurotoxins

Physiological Effect

1

Tetrodotoxin Saxitoxin μ-Conotoxins

Pore block

2

Veratridine Batrachotoxin Aconitine Grayanotoxins

Persistent activation Depolarization of resting potential Repetitive nerve firing

3

α-Scorpion toxins Sea anemone toxins δ-Astracotoxins

Prolonged channel opening

4

β-Scorpion toxins

Shifts in activation gating Repetitive nerve firing

5

Brevetoxins Ciguatoxins

Shifts in activation gating

6

δ-Conotoxins

Prolonged channel opening

7

Pyrethrins

Persistent activation Depolarization of resting potential Repetitive nerve firing

Sites 1-6 after Catterall et al. (4); Site 7 assigned arbitrarily as distinct from Sites 1-6.

With the exception of compounds acting at Site 1, all of the neurotoxins listed in Table 1 modify VGSCs to increase the probability or persistence of channel opening; the resulting enhanced influx of sodium ions causes neuronal excitation and results in aberrant neural signalling. These effects are particularly powerful because they disrupt nerve function at neurotoxin concentrations that modify only a small fraction of the available sodium channels; these toxins are therefore much more potent that would be expected based on conventional indices of pharmacological potency, which are based on concentrations producing half-maximal effects.

Sodium Channel-Directed Insecticides: Past and Present Pyrethrum The initial "discovery" of the VGSC as an insecticide target site occurred more than two centuries ago with the observation that dried pyrethrum flowers (Tanacetum cinerareaefolium) killed insects (6). In the 19th century insecticidal pyrethrum powder was widely available and used in Europe and Asia and was 39

imported into the United States as early as 1860. During the early decades of the 20th century pyrethrum (an organic extract of pyrethrum flowers) was produced commercially and widely used for insect control despite its expense and lack of environmental persistence. The introduction of more stable and less expensive synthetic insecticides limited the use of pyrethrum-based products, but pyrethrum is still produced commercially, registered for use, and remains an valuable insecticide option where low residual activity is important as well as in organic production. Pyrethrum was used widely as an insecticide long before the structures of its insecticidal constituents were known. The iterative application of newly-available methods and tools in natural products chemistry eventually identified six structurally-related esters, collectively called the pyrethrins (Figure 1) (7). Of these, pyrethrins I and II are the most abundant and typically exhibit the highest insecticidal activity. The extremely rapid action of pyrethrum on insects pointed to an effect on the nervous system. Early electrophysiological studies of the action of pyrethrins on invertebrate neurons using extracellular electrodes identified a consistent pattern of initial hyperexcitation, characterized by bursts of spontaneous and evoked action potentials, followed by nerve block (8). There is little further information on the mode of action of pyrethrins, either as a mixture of esters or as individual components. Therefore, much of what we infer about the action of pyrethrins is based on more detailed studies with allethrin, a close structural analog of pyrethrin I (Figure 2). Detailed voltage-clamp studies in both invertebrate and vertebrate axon preparations showed that allethrin modifies nerve membrane sodium currents by prolonging the opening of voltage-gated sodium channels. These findings established the voltage-gated sodium channel as the principal target site not only for allethrin but also, by inference, for the pyrethrins (8).

Figure 1. Structures and nomenclature of the pyrethrins. 40

Figure 2. S-bioallethrin.

DDT From our 21st century perspective it is difficult to appreciate the magnitude of the revolution in insect control afforded by DDT. Before DDT, chemical control of insect pests depended on inorganic compounds and natural products of botanical origin (including pyrethrum) (9). All of these agents were limited in effectiveness by their cost, performance or availability. DDT provided, for the first time in human history, relatively cheap and highly effective control of a broad spectrum of insect pests and disease vectors. The legacy of DDT, viewed from today’s perspective, is ambiguous (10). DDT not only protected crops but also saved lives; its use by the World Health Organization to control the mosquito vectors of malaria came very close to eradicating this disease before its use was curtailed. The success of DDT stimulated the search for new synthetic insecticides, which ultimately resulted in the array of highly effective insect control agents that are available today. However, the success of DDT also contributed to its downfall. In the 1950s and 1960s more than a billion tons of DDT were used in U.S. agriculture. The selection for resistance in populations of major pests led to increased use rates. The environmental persistence of DDT, initially viewed as a benefit allowing efficient control of insects, led to bioaccumulation in the environment and effects on nontarget species. In this way DDT contributed to the birth of the environmental movement and the creation of the U.S. Environmental Protection Agency, which in turned banned all uses of DDT in 1972. From the perspective of this review, the most significant aspect of DDT’s success as an insecticide lies in its validation of the voltage-gated sodium channel as a premier target for insect control agents (11). Early studies of the action of DDT on invertebrate neurons identified effects that were qualitatively similar to those of pyrethrins, and subsequent voltage-clamp experiments showed that DDT and allethrin modified sodium currents in nerve axons in a similar manner. The discovery of house fly strains resistant to both DDT and the rapid paralytic effects of pyrethrins provided further evidence for a shared mode of action. In the past two decades, genetic and molecular analyses of knockdown resistance in flies and other invertebrates have provided convincing evidence that mutations in genes encoding sodium channel α subunits underlie this type of resistance (12–14). 41

Pyrethroids The synthetic pyrethroids were developed over several decades by the iterative replacement of structural elements of the natural pyrethrins with other elements that preserved the steric and electronic features of the parent molecules (15, 16). The first breakthrough from this effort was the discovery of resmethrin (Figure 3) (17). Resmethrin surpassed both the pyrethrins and earlier synthetic analogs (e.g., allethrin; Figure 2) in both insecticidal activity and safety to mammals, but it remained susceptible to photochemical degradation and was therefore just as unstable in light and air as the pyrethrins and earlier synthetic compounds. Efforts to retain the favorable insecticidal and toxicological properties of resmethrin but improve environmental stability sufficiently to permit use in agricultural applications led, in the early 1970s, to the discovery of permethrin, deltamethrin and fenvalerate (Figure 3), the first pyrethroids developed for agricultural use (18). By 1995, more than 20 synthetic pyrethroids had been developed and registered for use worldwide (9).

Figure 3. Structures of resmethrin, permethrin, deltamethrin and fenvalerate. Pyrethroids proved to be surprisingly durable despite the impact of resistance in many important pest populations and the subsequent introduction of competing insect control technologies (Figure 4). Prior to the discovery of permethrin, deltamethrin and fenvalerate the commercial use of pyrethroids was insignificant, but by the late 1980s use of these compounds had grown to represent nearly 20% of the world insecticide market (19, 20). The introduction of commercial varieties of transgenic Bt cotton in the mid-1990s marked the beginning of a decline in pyrethroid use as transgenic cotton reduced the use of pyrethroids on that crop. Nevertheless, in 2013 still represented 17% of total insecticide sales worldwide (21). Pyrethroids disrupt nerve function by binding to a receptor site (Table 1, Site 7) that was first identified by the action of pyrethrins and subsequently also by the action of DDT (22). Different structural classes of pyrethroids produce qualitatively different effects depending on the duration of the pyrethroid-modified open state. Pyrethroids may also bind preferentially to either the resting (closed) 42

or the open state of the channel, depending on the pyrethroid and sodium channel preparation examined. The identification of pyrethroid resistance-associated mutations sodium channel genes and the functional characterization of wildtype and mutated channels in vitro have defined sodium channel domains that are determinants of pyrethroid sensitivity. More recently, this information has been employed to construct high-resolution structural models of the interaction of pyrethroid insecticides and DDT with these domains (12).

Figure 4. Pyrethroid use, measured as percent of the U.S. dollar value of worldwide sales, from 1972 to 2013 (19–21).

Sodium Channel Inhibitors Research at Philips-Duphar in the early 1970s identified novel substituted pyrazolines (PH-60-41, PH-60-42; Figure 5) with excellent insecticidal activity and signs of intoxication consistent with an action on the nervous system (23). Poor photostability and high soil persistence prevented the development of commercial products from this series. More than a decade later, Rohm and Haas reported the discovery of a second generation of pyrazoline-derived insecticides, exemplified by RH3421 (Figure 5) (24). These compounds retained the excellent insecticidal activity of the earlier pyrazolines but exhibited improved photostability and reduced environmental persistence. 43

Figure 5. Structures of early pyrazoline-based insecticides.

The development of commercial insecticide products from the pyrazoline series was ultimately halted by their high mammalian toxicity (25). Although the single-dose oral acute toxicity of these compounds was very low, longer-term dietary feeding studies revealed a delayed-onset acute neurotoxic response that occurred at doses lower than those producing acute intoxication following a single oral dose. Research to modify the core ring structure of the pyrazolines eventually yielded the first commercial insecticides from this group. Indoxacarb, an oxadiazine-based compound (Figure 6), was the first insecticide derived from the pyrazoline series to achieve commercial registration (26). Indoxacarb is a proinsecticide that is selectively hydrolyzed in insects to yield the ultimate toxicant, DCJW (Figure 6). The selective bioactivation of indoxacarb underlies its favorable selective toxicity and overcomes the toxicological barriers that were inherent in the pyrazoline series. Complete replacement of the central nitrogen-containing ring structure of the pyrazolines and the indoxacarb series led to the discovery of metaflumizone (Figure 6), the most recently-commercialized compound of this class (27).

Figure 6. Structures of indoxacarb, its bioactivation product DCJW, and metaflumizone. 44

Compounds identified as sodium channel inhibitors (SCIs; also called sodium channel blockers), though structurally diverse, are unified by a conserved core structure (Figure 7) (27) and a common mode of action on voltage-gated sodium channels (28). SCIs bind preferentially to sodium channels in the slow-inactivated state, a non-conducting conformational state that results from prolonged or repetitive depolarization (Figure 8). This interaction is very stable, resulting in the formation of a pool of insecticide-bound nonconducting channels and reducing the number of resting channels available for activation. The progressive sequestration of channels in the insecticide-stabilized slow-inactivated state eventually results in nerve block.

Figure 7. A proposed common core structure of SCI insecticides.

Figure 8. Conceptual model of the state-dependent action of SCI insecticides, illustrating the interconversion of channels in the closed (C),open (O), fast-inactivated (Ifast), and slow-inactivated (Islow) states. Site-directed mutagenesis studies showed that the SCI binding domain is likely to exist at the inner pore of the sodium channel protein, where it shares common molecular determinants with the binding site for local anesthetic and antiarrhythmic drugs (28). This site is distinct from the seven neurotoxin-binding domains on the channel protein (Table 1), and therefore represents a novel site and mechanism of insecticide action on sodium channels distinct from those identified for pyrethrins, pyrethroids and DDT. 45

Current Status of Sodium Channel-Directed Insecticides Figure 9 summarizes the relative importance, measured as worldwide sales, of different insecticide classes as reported by Sparks in 2013 (21). Compounds acting on ion channel targets accounted for approximately two-thirds of the insecticide market, with sodium channel-directed compounds (pyrethroids and SCIs) representing 19% of sales. Going forward, it is not clear whether the relative importance of sodium channel-directed insecticides will increase, remain stable or decrease. Pyrethroid use appears to be in a slow downward trend (see Figure 4) and the two commercially-available SCIs represent only a small segment of the market. However, the current regulatory pressure on neonicotinoid insecticide use may affect the future importance not only of sodium channel-directed compounds but also compounds acting at other targets.

Figure 9. Insecticide use by chemical class and target site, measured as total sales reported in 2013.

Future Sodium Channel-Directed Insecticides? The extensive exploitation of VGSCs for insect control by pyrethrum, DDT, and pyrethroids, together with widespread selection for target site-mediated resistance, might suggest that there is little future for the development of new insecticides acting at this target. However, the central importance of VGSCs in the processes underlying undesirable or damaging insect behaviors, the success of past and current insecticides acting at this target, and the rich pharmacology of the VGSC (Table 1) all argue for a role of VGSCs in future insect control, either by targeting currently-unexploited sodium channel binding sites or by the discovery of new target domains on the sodium channel protein. 46

Targeting Unexploited Binding Sites Of the seven discrete, well-characterized neurotoxin binding domains on the VGSC α subunit (Table 1), only one (Site 7) has been heavily exploited for insect control. Insecticidal agents acting at other receptor domains are likely to be unaffected by the domain-specific resistance limiting the action of insecticides acting at Site 7. The potential for this approach is illustrated by the discovery of novel isobutylamide insecticides that act at VGSC Site 2. The medicinal and insecticidal properties of naturally-occurring unsaturated aliphatic isobutylamides have been known since the early 19th century (29). Pellitorine (Figure 10), the earliest well-characterized example of this class, is the insecticidal principle isolated from the roots of Anacyclus pyrethrum. Pipercide (Figure 10), isolated from Piper nigrum, exemplifies naturally-occurring isobutylamides with extended aliphatic chains and aromatic substituents that exhibit greater insecticidal activity than pellitorine. Synthetic optimization from these natural product templates yielded compounds (e.g., BTG 502, Figure 10) with improved insecticidal activity (30). Moreover, BTG 502 was more effective against housefly strains carrying the super-kdr trait that confers high levels of pyrethroid resistance than against wildtype strains.

Figure 10. Structures of natural and synthetic isobutylamide insecticides.

47

In electrophysiological studies with insect nerve preparations, insecticidal isobutylamides induced repetitive activity followed by conduction block in housefly nerves, induced persistent sodium tail currents in cultured locust neurons, and inhibited the veratridine-dependent release of acetylcholine from cockroach presynaptic nerve terminals. These results pointed to an action on VGSCs that was qualitatively similar to that of pyrethroids but did not identify which sodium channel binding domain served as the isobutylamide target. More detailed pharmacological studies using mouse brain preparations and assays of radiosodium uptake and radioligand binding showed that BTG 502 and related insecticidal compounds acted as partial agonists at VGSC Site 2, thus identifying a novel insecticide target domain on the VGSC. Analysis of the pharmacological impact of point mutations introduced into insect sodium channels suggest that the binding site for pyrethroids and BTG 502 lie in close proximity to each other (31). Insecticidal isobutylamides have so far proved to be too unstable in the environment for commercial development. Nevertheless these compounds illustrate the potential for the discovery of new insecticides acting at sodium channel domains other than Site 7 that are not affected by resistance mechanisms that selectively alter the sensitivity of insect VGSCs to pyrethroids.

Discovering New Binding Sites The list of neurotoxin receptor sites on the VGSC (Table 1) is not finite. The discovery and exploitation of the local anesthetic receptor domain, first by drugs and more recently by SCI insecticides, illustrate the potential for the discovery of new chemistry that targets VGSCs by binding to previously-unrecognized receptor domains on the large VGSC protein. The most straightforward way to search for new agents that act at the VGSC would be to screen chemical libraries against insect VGSCs in vitro, specifically looking for compounds that function as sodium channel activators and are not affected by mutations that reduce the sensitivity of channels to pyrethroids. It would also be possible to counter-screen initial leads against human sodium channels in vitro to identify compounds with intrinsic selective toxicity to insects. Although attractive in principle there are two significant challenges inherent in this approach. First, to date insect sodium channels have been expressed in vitro only in the Xenopus laevis oocyte system (3). Although the automated high-throughput screening of channels expressed in oocytes is feasible, it remains more cumbersome that the use of stably-transformed cell lines as screening substrates. Further, we now know that the cellular environment of the oocyte expresses VGSCs with different functional and pharmacological properties than channels in cell lines or native neurons. For example, the actions of pyrethroids on rat Nav1.6 sodium channel complexes expressed in oocytes differ markedly from the actions of the same compounds on the identical channel complexes expressed in stably-transformed HEK293 cells when assayed under comparable voltage clamp conditions (32). Moreover, the properties of channels expressed in HEK293 cells, but not in oocytes, are similar to the functional and pharmacological properties of VGSCs expressed in mammalian neurons. 48

Therefore, in vitro screens for novel agents acting on insect VGSCs would benefit greatly from the development of stable cell lines expressing insect VGSCs. The second challenge to the in vitro screening approach is more generic. Intrinsic activity against a desirable target in vitro may be difficult to translate into compounds that can provide effective insect control in the real world and also exhibit environmental and toxicological profiles that will permit their development. These challenges are well illustrated by the current range of insecticidal compounds that act on sodium channels, all of which were identified by conventional screens for insecticidal action. The development or use of these compounds has been impeded or prevented by unacceptable persistence (either too short or too long) in the environment (e.g., pyrethrins, DDT, SCIs, isobutylamides) or toxicity to nontarget organisms (e.g., DDT, SCIs). In some cases these impediments were overcome to yield successful insecticides, but in other cases they prevented either continued use or commercial development. It is more likely that new insecticides that act at previously-unknown receptor domains on the VGSC will be identified the old-fashioned way – by empirical screening for insecticidal activity, followed by determination of mechanism of action. The essential function and the rich pharmacology of the VGSC suggest that screens for compounds that rapidly and irreversibly alter undesirable behaviors in pest insects will continue to identify compounds that act at the VGSC, and some of these compounds will likely be found to define new binding domains on the channel protein. The advances of the past two decades in the understanding the structure, function, and pharmacology of VGSCs at the molecular level will accelerate the further development of these novel compounds.

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J. E., Quistad, G. B., Eds.; Oxford University Press: New York, 1995; pp 217−233. Casida, J. E.; Quistad, G. B. Golden age of insecticide research: past, present or future? Annu. Rev. Entomol. 1998, 43, 1–16. Metcalf, R. L. A century of DDT. J. Agric. Food Chem. 1973, 21, 511–519. Davies, T. G. E.; Field, L. M.; Usherwood, P. N. R.; Williamson, M. S. DDT, pyrethrins, pyrethroids and insect sodium channels. IUBMB Life 2007, 59, 151–162. Zhorov, B.; Dong, K. Elucidation of pyrethroid and DDT receptor sites in the voltage-gated sodium channel. NeuroToxicology 2017, 60, 171–177. Soderlund, D. M.; Knipple, D. C. The molecular biology of knockdown resistance to pyrethroid insecticides. Insect Biochem. Mol. Biol. 2003, 33, 563–577. Rinkevich, F. D.; Du, Y.; Dong, K. Diversity and convergence of sodium channel mutations involving resistance to pyrethroids. Pestic. Biochem. Physiol. 2013, 106, 93–100. Elliott, M. Structural requirements for pyrethrin-like activity. Chem. Ind. 1969, 14, 776–781. Elliott, M. The relationship between the structure and the activity of pyrethroids. Bull. World Health Org. 1971, 44, 315–324. Soderlund, D. M. Resmethrin, the first modern pyrethroid insecticide. Pest Manage. Sci. 2014, 71, 801–807. Elliott, M. Established pyrethroid insecticides. Pestic. Sci. 1980, 11, 119–128. Lawrence, D. K.; Dunbar, S. J. Opportunities for neurotoxic compounds as crop protectants. In Progress in Neuropharmacology and Neurotoxicology of Pesticides and Drugs; Beadle, D. J., Ed.; Royal Society of Chemistry: Cambridge, 1999; pp 5−18. Pickett, J. A. New opportunities in neuroscience, but a great danger that some may be lost. In Neurotox’ 03: Neurotoxicological Targets from Functional Genomics and Proteomics; Beadle, D. J., Mellor, I. R., Usherwood, P. N. R., Eds.; Society of Chemical Industry: London, 2004; pp 1−10. Sparks, T. C. Insecticide discovery: evaluation and analysis. Pestic. Biochem. Physiol. 2013, 107, 8–17. Soderlund, D. M. Molecular mechanisms of pyrethroid neurotoxicity: recent advances. Arch. Toxicol. 2012, 86, 366–374. Mulder, R.; Wellinga, K.; van Daalen, J. J. A new class of insecticides. Naturwissenschaften 1975, 62, 531–532. Jacobson, R. M., A new class of insecticidal dihydropyrazoles. In Recent Advances in the Chemistry of Insect Control II; Crombie, L., Ed.; Royal Society of Chemistry: Cambridge, 1990; pp 206−212. Meier, G. A.; Silverman, R.; Ray, P. S.; Cullen, T. G.; Ali, S. F.; Marek, F. L.; Webster, C. A. Insecticidal dihydropyrazoles with reduced lipophilicity. In Synthesis and Chemistry of Agrochemicals II; Baker, D. R., Fenyes, J. G., Steffens, J. J., Eds.; American Chemical Society: Washington, DC, 1992; pp 313−326. 50

26. McCann, S. F.; Annis, G. D.; Shapiro, R.; Piotrowski, D. W.; Lahm, G. P.; Long, J. K.; Lee, K. C.; Hughes, M. J.; Myers, G. J.; Griswold, S. M.; Reeves, B. W.; March, R. W.; Sharpe, P. L.; Lowder, P.; Barnette, W. E.; Wing, K. D. The discovery of indoxacarb: oxadiazines as a new class of pyrazoline-type insecticide. Pest Manage. Sci. 2001, 57, 153–164. 27. Takagi, K.; Hamaguchi, H.; Nishimatsu, T.; Konno, T. Discovery of metaflumizone, a novel semicarbazone insecticide. Vet. Parasitol. 2007, 150, 177–181. 28. von Stein, R. T.; Silver, K. S.; Soderlund, D. M. Indoxacarb, metaflumizone, and other sodium channel inhibitor insecticides: mechanism and site of action on mammalian voltage-gated sodium channels. Pestic. Biochem. Physiol. 2013, 106, 101–112. 29. Jacobson, M. The unsaturated isolbutylamides. In Naturally-Occuring Insecticides; Jacobson, M., Crosby, D. G., Eds.; Marcel Dekker: New York, 1971; pp 137−176. 30. Elliott, M.; Farnham, A. W.; Janes, N. F.; Johnson, D. M.; Pulman, D. A.; Sawicki, R. M. Insecticideal amides with selective potency against a resistant (super-kdr) strain of houseflies (Musca domestica L.). Agric. Biol. Chem. 1986, 50, 1347–1349. 31. Du, Y.; Garden, D.; Khambay, B.; Zhorov, B.; Dong, K. Batrachotoxin,; pyrethroids and BTG 502 share overlapping binding sites on insect sodium channels. Mol. Pharmacol. 2011, 80, 426–433. 32. Soderlund, D. M.; Tan, J.; He, B. Functional reconstitution of rat Nav1.6 sodium channels in vitro for studies of pyrethroid action. NeuroToxicology 2017, 60, 142–149.

51

Chapter 4

Realizing the Potential: Improving a Microtransplantation Assay Based on Neurolemma-Injected Xenopus Oocytes An Ex Vivo Approach To Study Ion Channels in Their Native State Steven B. Symington,1 Edwin Murenzi,2 Abigail C. Toltin,1 David Lansky,3 and J. Marshall Clark*,2,4 1Department

of Biology and Biomedical Sciences, Salve Regina University, Newport, Rhode Island 02850, United States 2Molecular and Cellular Biology Program, University of Massachusetts, Amherst, Massachusetts 01003, United States 3Precision Bioassay Inc., Burlington, Vermont 05401, United States 4Department of Veterinary and Animal Sciences, University of Massachusetts, Amherst, Massachusetts 01003, United States *E-mail: [email protected].

Microtransplantation of rat brain neurolemma into the plasma membrane of Xenopus laevis oocytes is an ex vivo method used to study channels and receptors in their native state using standard electrophysiological approaches. Here we show that oocytes injected with adult rat brain neurolemma elicited ion currents upon membrane depolarization, which were increased by DDT and pyrethroid insecticides. Neurolemma incorporation and oocyte health varied, however, limiting the usefulness of the assay. A collection of changes to the assay procedure, data acceptance criteria, and analysis method yield substantially improved precision and hence, assay performance.

© 2017 American Chemical Society

Microtransplantation of Rat Brain Tissue into Oocytes To Examine Native Ion Channels Traditional electrophysiological approaches are powerful tools to examine the adverse effects of both natural toxins and synthetic toxicants on ion channels. Approaches used to study various ion channels include external cell recordings, whole cell patch clamp and heterologous expression of cloned channels, such as cRNAs injected into Xenopus oocytes (1–3). The speed of electronic data collection matches that of the gating processes of endogenous ion channels and allows a direct assessment of the effects of drugs and neurotoxicants on physiologically germane channel kinetic events. Nevertheless, these techniques are usually not amenable to high throughput screening and the physiological relevance of cRNA-injected oocytes has been questioned. For example, cRNA based approaches do not usually include multiple isoforms, variation in the level of expression, variations in channel activity due to axillary regulatory proteins, exon splicing or post-translational modifications, all of which occur in vivo. Alternatively, functional biochemical data collected from nervous system tissues from in situ preparations, such as tissue slices and isolated presynaptic nerve terminals (synaptosomes) are easily obtained from the mammalian CNS and retain a variety of functional attributes for study, such as ion flux, membrane depolarization and neurotransmitter release. However, these systems utilize non-physiological means to evoke depolarizing conditions and data collection occurs over time intervals in excess of those occurring in the intact nervous system (4). The model system evaluated here combines the strengths of both electrophysiological and biochemical techniques and reduces or eliminates many of the limitations mentioned above. This technique, pioneered by Ricardo Miledi, supports incorporation of neuronal tissue or cell lines expressing ion channels from a variety of brain regions and species, including humans (Figure 1) into Xenopus oocytes (5–12). The incorporation of intact lipid rafts from rat brain (neurolemma fragments), which are integrated into the plasma membrane of Xenopus oocytes, yields functional native voltage-sensitive sodium channels (VSSC) and auxiliary regulatory proteins in what is arguably a more physiologically-relevant ex vivo system to evaluate the effects of neurotoxic compounds, such as DDT and pyrethroids (13). This approach allows intact lipid membrane fragments along with associated target proteins to be microtransplanted into and function in the plasma membrane of the oocyte (Figure 1). The advantages of such an approach are that: a) receptors are in their native state within a lipid environment that mimics that found in the intact brain; b) a wide variety of voltage- and ligand-gated ion channels, transporters, and pumps are simultaneously available for study; c) rapid assessment of currents post injection (1-2 hours post-injection) is practical; d) small amounts of tissue are required; e) the procedure works with fresh or frozen tissue; f) the method can be used to the study differences among species, ages or tissue/organ location; g) selective currents can be separated pharmacologically and/or electrophysiologically and h) the method is amenable 54

to rapid data acquisition using two electrode voltage clamp (TEVC) techniques in a high-throughput format (e.g., Roboocyte2® system) (14).

Figure 1. Overview of experimental procedure used to analyze the effects of insecticides on rat brain neurolemma tissue microtransplanted into Xenopus laevis oocytes. (A) Roboinject (B) Robocyte with perfusion system (14).

Disadvantages include that: a) the approach is relatively uncharacterized; b) complex currents can result and; c) there is variability in the level of and orientation of the neurolemma incorporated into the oocyte plasma membrane. Results published on the incorporation of acetylcholine receptors from Torpedo marmorata electric organs showed variability in the amplitude of responses on an oocyte-to-oocyte basis as well the magnitude of the effect in the oocytes expressing the necessary proteins (8–15). Such differences could be due to variation among neurolemma preparations, variation in oocyte age or health associated with parents, variation in injection technique, neurolemma orientation, or any combination of these. 55

In the research presented herein, we evaluate the utility of using Xenopus oocytes, which are injected with rat brain neurolemma, to characterize the action of three well-understood neurotoxic insecticides, DDT and two pyrethroids: deltamethrin and permethrin; all three share a common mode of action on mammalian VSSC. Demonstrating that these insecticides function at VSSCs in this assay system would support use of this assay system as a useful ex vivo approach. This paper describes the improvements made to the assay methods, such as assay protocols for improving oocyte health, the development of criteria for the selection of neurolemma-injected oocytes, and data normalization procedures that, in combination, substantially reduce the variability in this model system. With the modifications described, we demonstrate that this assay system is a useful ex vivo approach to characterize the mode/site of action of a variety of neuroactive chemicals with the sensitivity, reproducibility and precision necessary for regulatory toxicological evaluations.

Initial Neurolemma-Injected Oocyte Assay Isolating and Estimating a TTX-Sensitive Inward Current In previously published research (16), we investigated the utility of using Xenopus laevis oocytes injected with 90 day old (PND90) adult rat brain neurolemma as a toxicologically-relevant, high-throughput, electrophysiological approach for studying the action of neurotoxicants on native ion channels in an ex vivo assay (Figure 1). The resulting current following neurolemma microtransplantation and oocyte depolarization was a complex outward current (Figure 2A), a component of which was sensitive to tetrodotoxin (TTX), a specific blocker of many VSSCs (Figure 2A). This TTX-sensitive component of the outward current was abolished by replacing sodium chloride with choline chloride in the recording buffer (16). Additionally, with calcium-activated chloride channels blocked by niflumic acid (NFA), a TTX-sensitive ‘inward’ post-depolarizatoin current of approximately 200 nA was measurable and spontaneously inactivated over approximately 50 ms. Thus, the TTX-sensitive inward current evoked upon depolarization appeared to undergo channel activation and inactivation in a manner similar if not identical to that seen during heterologous expression of VSSC/beta subunit cRNAs following co-injection into oocytes (17–22). To generate the concentration-dependent response curves (CDRCs), rows of each assay plate were assigned to concentrations of insecticide with one neurolemma-injected oocytes in each well on a 96-well micotiter assay plate. The concentrations used ranged from 1x10-9 M to 1x10-6 M. Concentrations were assigned to rows as follows: Row B = 1x10-9 M, Row C = 5x10-9 M, Rox D = 1x10-8 M, Row E = 5x10-8 M, Row F = 1x10-7 M, Row G =5x10-7 M, Row H = 1x10-6 M. Each oocyte was exposed to insecticide for 5 minutes by perfusing them at a flowrate of 0.45 ml/min with using a roboflow perfusion system (Figure 2B). 56

Figure 2. Insecticide induced TTX-sensitive inward currents in rat brain neurolemma tissue microtransplanted into Xenopus laevis oocytes in the presence of NFA. (A) TTX-sensitive inward currents (right panel) are generated from the difference between the treatment trace and TTX-insensitive trace (left panel). (B) Sample plate setup for microtransplantation assay. Insecticides concentrations (1x10-9-1x10-6 M) were perfused using a roboflow perfusion system for 5 minutes at a flowrate of 0.45ml/min. Pyrethroids concentrations were administrered in rows B-H of the 96 wellplate each pyrethroid perfusion varied between asssay plates.

In order to determine the effects of the test insecticides (DDT, permethrin or deltamethrin) on TTX-sensitive inward current, neurolemma-injected oocytes were analyzed in the presence of NFA (16). TTX-sensitive inward currents were determined by subtracting the experimental traces in the presence of TTX from the individual (oocyte-specfic) total current, both in the presence of NFA (Figure 2A). TTX-sensitive area under the curve values (AUC, nA x 50 ms) were determined for individual inward current traces from each oocyte. Pseudo-replicate oocytes at 57

each concentration of pesticide in each row of the assay plate (Figure 2B) yielded a TTX-sensistive AUC value, which were then (for the initial calculation method) averaged and then normalized using the mean of the same-plate NFA-treated controls via:

The data used to produce a CDRC were calculated from the mean AUC +/- S.E. values from 3 or more different biological replicates, each consisting of neurolemma from a single cohort of adults rats paired with a preparation of oocytes. CDRC data were fitted to a sigmoidal concentration-dependent response curve (3 parameter logistic equation) using GraphPad Prism ver 6.0. The asymptote of maximum response, the curve shape (or slope), and EC50 values were estimated.

Confirmation of Expected Perfomance Effect of DDT on TTX-Sensitive Inward Current It is well established that DDT acts on VSSCs, slowing the processes of channel inactivation and deactivation (17, 18). This action gives rise to increased sodium ion influx, leading to neuronal membrane depolarizing and “negative after potentials”, which ultimately are associated with repetitive discharges in the nerve axon and a likely cause of the “DDT jitter” syndrome typical of DDT poisoning. This action is similar, if not identical, to the tremor or T-syndrome produced by Type I pyrethroids, such as permethrin (19–23). With the above in mind, we investigated the action of DDT on neurolemmainjected oocytes (16). The rationale for this approach was that if the action of DDT on the TTX-sensitive inward current measured from neurolemma-injected oocytes mimicked the action of DDT on heterologous expressed VSSCs, it would provide support of the toxicological relevancy of this ex vivo approach. Using the method described above, many of the issues/limitations of previously used approaches (e.g., heterologous expressed channels, non-neural tissues, biochemical assays) are reduced or eliminated, supporting more precise comparisons of the neurotoxicity of various chemicals under various conditions (i.e.; animal age). Using the microtransplantation approach to obtain CDRCs, DDT increased TTX-sensitive inward currents in a concentration-dependent manner by increasing Na+ influx during depolarization (Figure 3A and B) primarily by slowing the inactivation kinetics of VSSC (Figure 3C). The non-toxic DDT metabolite, DDE, however, resulted in no observable concentration-dependent response (16). These results indicate a clear structure-activity relationship for DDT versus DDE on the VSSC current that has been pharmacologically isolated with TTX and in the presence of NFA using rat brain neurolemma microtransplanted oocytes.

58

Figure 3. DDT increases TTX-sensitive current in rat brain neurolemma injected oocytes. (A) Neurolemma-injected oocytes in the presence of NFA and increasing concentrations of DDT. TTX-sensitive area-under the curve (AUC) values (nA x 50 ms) were determined from individual inward current traces during depolarization. (B) Concentration-dependent response curve illustrating the effect of DDT. Percent over control values were determined as follows: ((Treatment TTX-sensitive AUC – No treatment TTX-sensitive AUC)/No treatment TTX-sensitive AUC) x 100). (C) Inactivation tau values are increased in the presence of 10-6 M DDT. Adapted with permission from Murenzi et al., 2017 (16). The inactivation tau 0.5 (t0.5) values can be estimated from the current traces given in Figure 1C as 2.6 ms for untreated cells and 5.9 ms for DDT-treated cells, a 1.3-fold increase. In the present work, we determined the inactivation tau0.5 value in the presence of 1 µM DDT as 6.7 ms, a 1.5-fold increase compared to the 4.4 ms control value. These results are similar to those reported by Song et al. (24, 25), who report that 10 µM DDT slowed the inactivation kinetic of TTX-sensitive VSSCs in rat dorsal root ganglion cells following a 20 ms step depolarization. We conclude that this ex vivo assay is a toxicologically-relevant approach to examine native receptors in their endogenous states.

Effect of Pyrethroids on TTX-Sensitive Inward Current It is well also well established that pyrethroids modify VSSCs by slowing the inactivation and deactivation channel kinetic processes (26–28). This action also gives rise to increase sodium ion influx, leading to neuronal membrane depolarization, nervous system excitation, convulsions and death. The addition of increasing concentrations of permethrin (Figure 4A) or deltamethrin (Figure 4D) from 10-9 to 10-6 M resulted in an increase of TTX-sensitive inward current on VSSCs microtransplanted into Xenopus oocytes. Calculation of AUC values at each concentration and their normalization to percent over control values for each biological replicate using (1) allowed composite CDRCs to be similarly generated. Increasing concentrations of either permethrin (Figure 4B) or deltamethrin (Figure 4E) progressively increased the percent over control values. The increases in the percent over control values 59

were primarily due to the prolongation of inactivation process as seen at the 10-6 M concentration (Figure 4C for permethrin; Figure 4F for deltamethrin). These results illustrate that the pyrethroid effect on late current is similar to results previously obtained from other heterologous expression systems, supporting the notion that this ex vivo preparation is a toxicologically-relevant approach to examine native receptors in their endogenous states (13).

Figure 4. Effects of increasing concentrations of pyrethroids on depolarization-evoked, TTX-sensitive inward currents associated with rat brain neurolemma microtransplanted into Xenopus oocytes in the presence of NFA, a Ca2+-activated chloride channel blocker. A) Electrophysiological TTX-sensitive current traces illustrating the effects of increasing concentrations of permethrin. (B) Concentration-dependent response curve (CDRC) illustrating the effect of permethrin. (C) Effect of 10-6 M permethrin on the inactivation tau value. D) Electrophysiological TTX-sensitive current traces illustrating the effects of increasing concentrations of deltamethrin. (E) CDRC illustrating the effect of deltamethrin. (F) Effect of 10-6 M deltamethrin on the inactivation tau value. TTX-sensitive inward currents were determined by subtracting the experimental traces in the presence of TTX from the total current. Percent over control values were determined using equation 1. Inactivation tau values were obtained by fitting the late current traces to an exponential decay equation during inactivation using Origin (Ver 8.6, Origin Lab, Northampton, MA). An asterisk (*) indicates that the sample mean is significantly different from the mean of control treated (NFA) oocytes (one-sample Student’s t-test, 10-6 M: p95%), piracetam (>90%), pramiracetam (>98%), and rolipram (>98%). Quisqualic acid (>99) and aniracetam (>99%) were purchased from TOCRIS Bioscience (Ellisville, MO, USA). Phenylpiracetam (>98%) was purchased from Cayman Chemical Company (Ann Arbor, MI, USA). Propoxur (≥99%) was obtained from Fluka (Morris Plains, NJ, USA). The majority of these compounds are water soluble, and therefore, were dissolved in water or buffer, except aniracetam and cyclothiazide, which were dissolved in DMSO (final DMSO concentration was 0.1%).

Figure 1. Chemical structures of glutamatergic agonists, cyclothiazide, and racetams. 113

Insects Aedes aegypti mosquitoes were obtained from the United States Department of Agriculture – Agricultural Research Service, Center for Medical, Agricultural, and Veterinary Entomology (USDA-ARS, CMAVE), Gainesville, FL, USA as 3rd or 4th instar larvae, and reared as previously described (17). Briefly, the larvae were held in tap water and fed a diet that consisted of 3-parts liver powder (MP Biomedical, Solon, OH, USA) to 2-parts Brewer’s yeast (MP Biomedical). Adult mosquitoes were provided a cotton ball soaked with 10% sugar water for sustenance. Mosquitoes were maintained at 28°C, relative humidity >60%. Adult female mosquitoes that were 1-5 days post-emergent were used for feeding and injection studies. Oregon-R, a susceptible strain of Drosophila melanogaster has been maintained in culture at the University of Florida (Gainesville, FL, USA) since 2009. Flies were reared in plastic vials on artificial media that was purchased from Carolina Biological Supply (Burlington NC, USA). Adult females that were 1-2 weeks old were used for feeding experiments. Toxicity and Paralysis Bioassays Oral toxicity of glutamatergic compounds was assessed against adult female Ae. aegypti and D. melanogaster. Mosquitoes were anaesthetized on ice, transferred to a holding container, and starved for 6 hr. D. melanogaster were also starved for 6 hr and anaesthetized using carbon dioxide before being transferred to a test tube. Compounds were delivered by dissolving in 10% sugar water and 1 mL was applied to a cotton ball that stoppered a glass vial or test tube that held adult insects. Mortality of the test compounds was determined after 48 hr of exposure. A headless larval bioassay (18) was employed to examine the rapid paralytic activity against fourth-instar Ae. aegypti larvae, because pilot studies with L-aspartic acid showed it was inactive against intact larvae. Briefly, fourth-instar Ae. aegypti larvae were decapitated by pulling off the head using forceps. Decapitated larvae were placed into 5 mL of mosquito physiological saline, which was composed of (mM) sodium chloride (154), calcium chloride (1.4), potassium chloride (2.7), and HEPES (1.2); pH was adjusted to 6.9 (19). Motor responses to manual probing of the headless larvae were monitored for five hours post-decapitation. At this time, if the larva displayed little or no movement after being gently probed with a needle, it was classified as paralyzed. Control paralysis under identical conditions was 1 were L-BMAA, quisqualic acid, and willardiine (Table 1). The paralytic effect of the putative GluRd desensitization blockers (cyclothiazide and racetams) was also investigated against fourth-instar headless larvae (Table 2). Piracetam and pramiracetam had the lowest PC50 values ( 10 ng/mg, but ≤ 100 ng/mg. Finally, the least active compounds showed an LD50 value > 100 ng/mg. Five out of the nine-glutamatergic agonists were in the high activity category (LD50 ≤ 10 ng/mg) when injected into adult female Ae. aegypti. The five compounds were essentially equipotent, and included L-BMAA, quisqualic acid, domoic acid, kainic acid, and aspartic acid. The moderately active glutamatergic agonists were AMPA, willardiine, and NMDA. L-Glutamic acid was the least active glutamatergic agonist when injected into adult female Ae. aegypti (Table 3). 116

Table 2. Paralytic Activity of Cyclothiazide and Racetam Compounds against Headless Fourth-Instar Ae. aegypti Larvae Drug aniracetam cyclothiazide phenyl-piracetam piracetam pramiracetam rolipram

PC50, ppm1

Slope ± SE

χ2 value (df)

150 (74 – 400)

0.50 ± 0.08

7.98 (30)

34 (22-51)

1.24 ± 0.20

15.06 (15)

136 (101– 201)

2.28 ± 0.49

9.58 (19)

9 (6 – 13)

0.85 ± 0.09

13.73 (38)

10 (3-26)

0.87 ± 0.19

1.70 (5)

50 (36 – 72)

1.71 ± 0.24

11.21 (17)

Concentration that resulted in 50% larval paralysis in headless fourth-instar Ae. aegypti. Data are presented as the PC50 with the 95% confidence interval in parentheses, 5 hr postdecapitation.

1

Piracetam was the most toxic of the glutamatergic desensitization modulators when injected into adult mosquitoes. Moderately toxic compounds included rolipram and pramiracetam. Finally, aniracetam and phenyl-racetam were the least active (Table 4). Propoxur was again used as a benchmark for comparing relative toxicity, and had an injected LD50 of 0.14 (0.1-0.2) ng/mg (20). Propoxur was 29-fold more toxic than L-BMAA and piracetam, which were the most toxic glutamatergic agonist or modulator, respectively. The slope values for injected glutamatergic agonists/modulators into adult female mosquitoes were more steep (values greater than 2; Tables 3 and 4) when compared to headless larval slope values. A linear regression correlation analysis was performed between toxicities in the fourth-instar headless larval assay and by intrathoracic injection of adult female mosquitoes (Figure 2). When all glutamatergic and racetam compounds where included in linear regression, a correlation coefficient (R2) of 0.70 was obtained (dotted line, Figure 2). However, the R2 increased to 0.97 when kainic acid was excluded from the regression analysis, which had variable results between the two bioassays (solid line, Figure 2). 117

Table 3. Intrathoracic Injection and Oral Toxicity of Glutamatergic Compounds against Adult Female Ae. aegypti LD50 (ng/mg)1

Slope ± SE

χ2 value (df)

Oral toxicity2

AMPA

12 (9 – 19)

2.80 ± 0.57

0.88 (2)

50%

L-aspartic acid

7 (6 – 10)

3.26 ± 0.59

0.58 (2)

0%

L-BMAA

4 (3 – 5)

3.73 ± 0.79

0.11 (2)

61 (40-79)

domoic acid

5 (4 – 7)

3.28 ± 0.64

1.43 (2)

294 (167-340)

L-glutamic acid

371 (304 – 473)

2.63 ± 0.41

6.09 (10)

20%

kainic acid

7 (6 – 10)

3.26 ± 0.59

0.58 (2)

40%

98 (70 – 153)

2.45 ± 0.55

0.55 (2)

10%

quisqualic acid

4 (3 – 6)

3.19 ± 0.65

0.79 (2)

7%

willardiine

82 (61 – 116)

2.91 ± 0.62

0.46 (2)

20%

Drug

NMDA

1 Mortality 24 hr post-injection, given as LD 50 with 95% confidence intervals in parentheses. 2 Oral toxicity of glutamatergic compounds determined at 48 hr and given as the LC50 in ppm (95% confidence intervals) or the percentage of mortality at 1 mg/mL.

Feeding Bioassays and Aedes Aegypti and Drosophila melanogaster The oral toxicity of the glutamatergic compounds was investigated in adult female Ae. aegypti and D. melanogaster. Initially, the 48 hr toxicity of these compounds was measured at a high concentration (1 mg/mL) of the toxin delivered in 10% sugar water (w/v). D. melanogaster displayed little toxicity (≤ 30% mortality at 48 hr) for any glutamatergic compound, with the majority of the observed toxicities ranging from 0-10% (data not shown). For Ae. aegpyti, no or low mortality was observed for L-aspartic acid, NMDA, quisqualic acid, and willardiine at 1 mg/mL (mortality less than 50%; Table 3). For the remaining glutamatergic compounds tested L-BMAA was about 5-fold more active than domoic acid (Table 3). Three out of five racetams tested (piracetam, aniracetam, and phenyl-racetam) resulted in low mortality (< 30%) when tested at a high concentration of 1 mg/mL. Pramiracetam and rolipram had LC50 values of 389 and 314 ppm, respectively; however, they were still significantly less toxic than cyclothiazide (Table 4). For comparison, propoxur had an LC50 of 0.33 (0.27-0.38) ppm in this assay, making it 185-fold more toxic than L-BMAA. 118

Table 4. Intrathoracic Injection (LD50) and Oral Toxicity of Cyclothiazide and Racetam Compounds against Adult Female Ae. aegypti Drug

LD50, ng/mg1

Slope ± SE

χ2 value (df)

Ae. aegypti oral toxicity 2

aniracetam

142 (101 – 247)

2.61 ± 0.63

0.0073 (2)

10%

cyclothiazide

14 (11 – 21)

2.33 ± 0.35

8.38 (10)

120 (83-167)

phenylpiracetam

171 (120-255)

2.28 ± 0.49

2.43 (2)

30%

piracetam

4 (3 – 5)

2.96 ± 0.66

0.51 (2)

0%

pramiracetam

76 (54 – 118)

2.16 ± 0.46

1.44 (2)

389 (251 – 502)

rolipram

50 (38 – 67)

2.88 ± 0.56

0.34 (2)

314 (226 – 368)

LD50 with 95% confidence interval in parentheses measured at 24 hr post-injection. 2 Oral toxicity (48 hr) of test compounds represented as the LC50 in ppm (95% confidence interval) or the percentage of mortality at 1 mg/mL. 1

Figure 2. Correlation of Ae. aegypti headless fourth-instar larval assay (x-axis) versus the intrathoracic injection of adult Ae. aegypti female mosquitoes (y-axis). Inclusion of all 15 of the tested glutaminergic compounds yielded the dotted regression line, with R2=0.70. Exclusion of kainic acid (KA) gave the solid regression line, with R2=0.97. 119

Table 5. Paralytic Activity of Glutamatergic Compounds Mixed with 1 ppm Piracetam against Headless Fourth-Instar Ae. aegypti Larvae Calculated Additive PC50, ppm1

AR2

Actual PC50, ppm1

Actual SR3

AMPA

7 (2 – 16)

4.4

31 (14 – 72)

1.0

L-aspartic acid

3 (1 – 6)

4.0

10 (4 – 24)

1.2

L-BMAA

3 (1 – 7)

3.0

8 (2 – 18)

1.1

cyclothiazide

15 (8 – 25)

2.3

8 (3 – 20)

4.3

domoic acid

9 (3 – 26)

3.7

31 (11 – 83)

1.1

L-glutamic acid

96 (40 – 302)

3.2

164 (82 – 399)

1.9

kainic acid

89 (39 – 266)

2.3

42 (20 – 94)

4.9

NMDA

38 (17 – 87)

3.3

92 (45 – 211)

1.4

4 (1 – 8)

2.0

6 (3 – 13)

1.3

59 (41 – 86)

1.6

65 (35 – 135)

1.5

Compound

quisqualic acid willardiine

Calculated Additive and Actual PC50 values are given, with the 95% confidence interval in parentheses. 2 Additivity Ratio (AR) calculated by dividing the PC50 of the glutamatergic alone by the PC50 obtained when adding the 10% paralysis expected from piracetam (1 ppm). 3 Synergistic Ratio (SR) calculated by dividing the PC50 of the glutamatergic alone by the actual PC50 obtained when mixed with the glutamatergic with piracetam (1 ppm). 1

Synergism of Glutamatergic Compounds with Piracetam Since racetams reduce desensitization of mammalian iGluR (11–16), piracetam, the most active racetam in the headless larval assay, was further tested in this assay at 1 ppm in combination with other glutamatergic compounds to document any synergistic effects (Table 5). As part of the test for synergism, a theoretical additive PC50 value was calculated by probit analysis of the summed paralysis obtained with 1 ppm piracetam (10%) added to that obtained at each concentration of glutamatergic agonist alone, taken from Table 2. It was important to account for an additive effect of piracetam due to the low slope values observed in the headless larva assay (Table 1). 120

As described above, an additive effect of 1 ppm piracetam would be expected to decrease the PC50 of the agonists about 2-4 fold (Table 5) compared to that observed with agonist alone (Table 1). Cyclothiazide and kainic acid were the only two glutamatergic compounds tested where the SR was greater than the calculated AR, but given the extensive overlap in the 95% confidence limits, the difference was not statistically significant. SR values less than the AR indicate that the piracetam effect was less than additive, indicating some kind of antagonism was occurring with the majority of the glutamatergic agonists (Table 5).

Discussion The insect glutamatergic system plays an important role in the excitatory neuromuscular and central synaptic transmission, but it has not been a successful target of insecticide development. The lack of success at exploiting the GluRd has been attributed to a lack of chemistry that can penetrate the insect cuticle (21). In this chapter, we investigated the toxicity of known glutamatergic agonists and desensitization modulators in two mosquito life stages using bioassays that circumvented the cuticular barrier. In both headless larval and adult injection assays, fairly potent intrinsic activity of GluRd agonists was found. In the headless larval assay, we observed low slope values (< 1) in probit toxicity analysis (Tables 1 and 2). These shallow slopes suggest a slow rate of penetration of the test compounds through the cervical opening. As expected, the LD50 obtained by injecting test compounds into adult mosquitoes resulted in greater slope values and narrower 95% confidence limits. The high correlation coefficient for the toxicity data of headless larval and intrathoracic injection studies indicates a similar receptor type mediates toxicity in both life stages (Figure 2). The high potency of L-aspartic acid in larval and adult assays was unanticipated. This amino acid is known to stimulate insect muscle glutamate receptors, but usually requires higher concentrations than L-glutamate (22). However, it is worth noting that no information is available on relative potencies of these amino acids on mosquito receptors, nor their affinity for active transporters at either peripheral or central synapses that might impact toxicity. The low sensitivity of L-glutamic acid we observed in Ae. aegypti larvae and adults have previously been reported in other insects (21). When L-glutamic acid was injected into adult male Lucilia sericata, it resulted in an LD50 of 7800 ng/mg (23), which is much higher than the LD50 value we found in adult female Ae. aegypti (LD50 = 371 ng/mg). A previous review (24) indicated that variable concentrations of L-glutamic acid can be found in insect hemolymph and it discussed possible mechanisms that could serve to protect the neuromuscular junction from the circulating glutamic acid found in the hemolymph. Such mechanisms may include a diffusion barrier, which has been characterized as a connective tissue sheath surrounding muscle cells, as described in Schistocerca gregaria (6). This protective sheath acts as a barrier to glutamic acid, thereby protecting the nerve-muscle synapses. Additionally, glial cells and tracheal sheath cells may play a pivotal role in the uptake of glutamic acid at the neuromuscular junction, which has been described in cockroaches (24, 25). While barriers 121

might contribute to the low paralysis and toxicity we observed with L-glutamic acid in larval and adult mosquitoes, they would have to be permeable to other glutamatergic compounds having similar physical properties, suggesting that specific active transport is a more likely mechanism. The modest oral toxicity of glutamatergic compounds in Ae. aegypti and D. melanogaster (Tables 3 and 4) was disappointing. The most toxic compound via this route of administration was L-BMAA. Quisqualic (10) and domoic (9) acids were previously shown to inhibit peristaltic movements in the cockroach gut (10), but only domoic acid was orally toxic to Ae. aegypti in the present study. Overall, there was a species-specific oral toxicity of glutamatergic compounds favoring Ae. aegypti over D. melanogaster (Table 3 and 4). Additionally, L-BMAA is known to cause neurotoxicity from dietary exposure to this amino acid (26), and domoic acid ingestion is implicated as a cause of human amnesic shellfish poisoning (27). We find no reports on the injected toxicity of glutamatergic compounds into D. melanogaster to use for comparison with Ae. aegypti. However, domoic acid has been injected into adult male Peripleneta americana, resulting in an LD50 of 800 ng/insect, which was similar to allethrin (injected LD50 500 ng/insect) (9). Assuming that the average weight of an adult male P. americana is between 500-700 mg (weights obtained from 5 adult male cockroaches in our laboratory), domoic acid has an approximate LD50 between 1.1-1.6 ng/mg. This LD50 for domoic acid is not too dissimilar from that observed in adult female Ae. aegypti (5 ng/mg). Therefore, we hypothesize that species-selective toxicity may be induced by glutamatergic compounds, but its magnitude will differ by route of exposure. The racetam compounds (aniracetam, piracetam, pramiracetam, phenylpiracetam, and rolipram) are nootropics, cognitive enhancers that extend attention span and memory (28). Cyclothiazide (a diuretic and anti-hypertensive) and aniracetam have been reported to enhance glutamate-evoked currents, decrease receptor desensitization, and prolong synaptic current (14–16). Based on these previous findings, we hypothesized that piracetam could synergize or enhance the activity of glutamatergic agonists. Accordingly, synergism of glutamatergics by piracetam was examined in the headless larval assay at 1 ppm (Table 5). The lack of synergism observed between piracetam and agonists suggests that mosquito GluRd are pharmacologically different from mammalian iGluR, which may be beneficial in the future development of GluRd-selective insecticides. In conclusion, this study reports on the toxicity of glutamatergic agonists in Ae. aegypti. The insect GluRd is a physiological relevant receptor located in the central nervous system of insects, but also in the periphery, so it could serve as an excellent target for mosquito control. A drawback of the available lead compounds, some of which were studied in this chapter, is that they are highly polar and therefore have poor barrier penetration in insects. In addition, several of the compounds have deleterious effects in mammals from oral exposure, and selectivity would be an important factor in the development of any strategy for targeting the GluRd. The lack of synergism of racetams and agonists suggests further differences in the pharmacology of mammalian and insect iGluRs that will be the focus of future mode of action research.

122

Acknowledgments This work was supported by USDA Specific Cooperative Agreement 58-0208-5-001 to J.R.B. as part of the Deployed War Fighter Research Program. The authors are thankful to Dr. Dan Kline (USDA-ARS, CMAVE) and his insectary staff for providing the mosquitoes used in this study.

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15. Isaacson, J. S.; Nicoll, R. A. Aniracetam reduced glutamate receptor desensitization and slows the decay of fast excitatory synaptic currents in the hippocampus. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 10936–10940. 16. Mayer, M. L.; Partin, K. M.; Patneau, D. K.; Wong, L. A.; Vyklicky, L.; Benveniste, M.; Bowie, D. Desensitization at AMPA, kainate and NMDA receptors. In Excitatory Amino Acids and Synaptic Transmission, 2nd ed.; Wheal, H., Thomson, A., Eds.; Academic Press Inc.: San Diego, CA, 1995; pp 89−113. 17. Pridgeon, J. W.; Pereira, R. M.; Becnel, J. J.; Allan, S. A.; Clark, G. G.; Linthicum, K. J. Susceptibility of Aedes aegypti, Culex quinquefasciatus Say, and Anopheles quadrimaculatus Say to 19 pesticides with different modes of action. J. Med. Entomol. 2008, 45, 82–87. 18. Islam, R. M.; Bloomquist, J. R. A method for assessing chemically-induced paralysis in headless mosquito larvae. MethodsX 2015, 2, 19–23. 19. Hayes, R. O. Determination of a physiological saline for Aedes aegypti (L.). J. Econ. Entomol. 1953, 46, 624–627. 20. Larson, N. R.; Carlier, P. R.; Gross, A. D.; Islam, R. M.; Ma, M.; Sun, B.; Totrov, M. M.; Yadav, R.; Bloomquist, J. R. Toxicology of potassium channel-directed compounds in mosquitoes. NeuroToxicology 2017, 60, 214–223. 21. Miller, T. A. The insect neuromuscular system as a site of insecticide action. In Pesticide and Venom Neurotoxicity; Shankland, D. L., Hollingworth, R. M., Smith, T., Eds.; Plenum Press, New York, 1978; pp 102−111. 22. Usherwood, P. N. R.; Machili, P. Pharmacological properties of excitatory neuromuscular synapses in the locust. J. Exp. Biol. 1968, 49, 341–361. 23. Hart, R. J.; Potter, C.; Wilson, R. G. Factors governing the toxicity of putative synaptic transmitters and their analogues when injected into the haemocoel of adult male Lucilia sericata. Pestic. Sci. 1977, 8, 722–734. 24. Faeder, I. R.; Salpeter, M. M. Glutamate uptake by a stimulated insect nerve muscle preparation. J. Cell Biol. 1970, 46, 300–307. 25. Salpeter, M. M.; Faeder, I. R. The role of sheath cells in glutamate uptake by insect nerve muscle preparations. Prog. Brain Res. 1971, 34, 103–114. 26. Duncan, M. W.; Kopin, I. J.; Crowley, J. S.; Markey, S. P. Quantification of the putative neurotoxin 2-amino-3-(methylamino)propanoic acid (BMAA) in cycadales: analysis of the seeds of some members of the family Cycadaceae. J. Anal. Toxicol. 1989, 13, 169–175. 27. Teitelbaum, J. S.; Zatorre, R. J.; Carpenter, S.; Gendron, D.; Evans, A. C.; Gjedde, A.; Cashman, N. R. Neurologic squelae of domoic acid intoxication due to the ingestion of contaminated mussels. N. Engl. J. Med. 1990, 322, 1781–1787. 28. Vincent, G.; Verderese, A.; Gamzu, E. The effects of aniracetam (Ro 135057) on the enhancement and protection of memory. Ann. N. Y. Acad. Sci. 1985, 444, 489–491.

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Chapter 8

Metabolites of Induced Fungi: A Potential Chemical Library for Next-Generation Pesticides S. Furutani, M. Ihara, K. Kai, H. Hayashi, and K. Matsuda* Department of Applied Biological Chemistry, Faculty of Agriculture, Kindai University, 3327-204 Nakamachi, Nara 631-8505, Japan Graduate School of Life and Environmental Sciences, Osaka Prefecture University, Sakai, Osaka 599-8531, Japan *E-mail [email protected].

Although some fungi produce mycotoxins that render food on which they grow inedible, other fungi produce metabolites that, in addition, to human and animal healthcare, are useful in pest control. Penicillium simplicissimum AK-40 produces okaramines as an insecticide when grown on okara, a by-product of soybean curd production. Okaramines selectively activate glutamate-gated chloride channels expressed only in the nervous system of invertebrates. Asperparalines from Aspergillus japonicus JV-23 and chrodrimanins from Talaromyces sp. YO-2, both of which were isolated similarly to okaramines, selectively block insect nicotinic acetylcholine and γ-aminobutyric acid receptors, respectively. These results suggest that fungi induced by certain plant factors have potential for producing next-generation pesticide leads.

Introduction In the late 1980s, Dr. Hideo Hayashi, now Emeritus Professor of Osaka Prefecture University, began to explore environmentally benign, next-generation pesticide leads with new skeletons from fungal metabolites due to their diverse metabolite structures (1). Based on literature, potato-dextrose and Czapek-Dox © 2017 American Chemical Society

culture media etc. were found to be suitable for proliferating fungi. However, he did not employ known media but instead used okara, a by-product of soybean curd (tofu) production, since it was very cost-effective, and its use was still unknown. He screened various fungal metabolites for toxicity using larvae of Bombyx mori and discovered two indole alkaloids, okaramine A and B, from the metabolites of Penicillium simplicissimum AK-40 (Figure 1) (2), which act as insecticides with LD50 (dose that kills 50% of insects) values of 8 and 0.2 μg/g diet, respectively. Both compounds possess a unique eight-membered azocine ring structure. Additionally, a searh for other okaramines and was able to further identify 16 okaramines (1). Okaramine B possesses a methoxy and a hydroxy group as well as a four-membered azetidine ring in addition to the azocine and indole rings and shows the highest insecticidal activity against the silkworm larvae among the 18 okaramines (1). Partial hydrogenation of its azocine ring led to reduced activity (3), and okaramine C (4) lacking the azocine ring was less potent than okaramine B. These results indicate the essential role of the azocine ring in okaramine potency.

Figure 1. Insect–active metabolites produced by fungi in okara.

Motivated by the discovery of okaramines, Dr. Hayashi continued to explore insect-active fungal metabolites with distint skeletons from okaramines (1). During this process, meroterpenoids, such as chrodrimanins, as well as other classes of indole alkaloids, such as asperparalines, were discovered (Figure 1) (1). When tested by oral application, chrodrimanin B (5) and E (6) show the highest insecticidal activity among the chrodrimanins tested on the silkworm larvae with an LD50 value of 10 μg/g diet, while asperparaline A induces paralysis at the same dose (7). Their unique and complex structures inspired total synthesis (8–11). However, their mechanism of action remains unknown, and thus the authors have worked to resolve their enigma. 126

Figure 2. Selective blocking action of asperparaline A on ACh-induced currents in the silkworm larval neurons. Asperparaline A was applied for 1 min and then co-applied with 10 μM ACh (left), 30 μM GABA (center) and 30 μM L-glutamate (right). Applications of ACh, GABA and L-glutamate are indicated by horizondal lines, while those of asperparaline are indicated by dashed horizontal lines. Adapted with permission from ref. (12). Copyright 2011 PLoS.

Establishing a Patch-Clamp Recording Technique for Silkworm Larval Neurons and Clarifying the Mode of Action of Asperparaline All the fungal products shown in Figure 1 exhibited toxicity with a rapid onset of action in the silkworm larvae, indicating possible interactions with the larval nervous system (1). Therefore, we employed patch-clamp electrophysiology to investigate their actions on the ion channels expressed in the silkworm larval neuron. When applied alone, asperparaline A did not induce any currents in the neurons. However, it blocked the responses to acetylcholine (ACh) when co-applied with ACh, while scarcely influencing the responses to γ-aminobutyric acid (GABA) and L-glutamate (Figure 2) (12). The IC50s, half-maximal inhibitory concentrations, of asperparaline A for the peak and steady-state ACh-induced currents were 20 and 40 nM, respectively. By contrast, 10 μM asperparaline A only slightly attenuated the peak current amplitude of the response to ACh of the avian α4β2, α3β4 and α7 nAChRs expressed in Xenopus laevis oocytes (12), suggesting a high selectivity for insect nAChRs.

Resolving the Enigma of Okaramines Having confirmed the effectiveness of the patch-clamp electrophysiology in elucidating the mode of action of asperparaline A, we next investigated the mechanism of action of okaramines. Unlike asperparaline A, okaramine B was capable of inducing inward currents in a concentration-dependent manner. In the current-voltage relationship, the peak current amplitude of the okaramine-induced current crossed the X-axis at a potential close to the equilibrium potential for the Cl- ion. Furthermore, the reversal potential shifted to a more positive potential when the extracellular Cl- ion concentration was reduced, suggesting that the okaramine-evoked currents were mediated by Cl- ions. Fipronil, a blocker for 127

ligand-gated chloride channels, attenuated the peak response of okaramine B, whereas mechamylamine, an antagonist of nicotinic acetylcholine receptors, had no such effect, suggesting that interactions at the ligand-gated chloride channels dictate the reversal potential for the okaramine-induced current (13). Glutamate-gated chloride channels (GluCls) as well as GABA-gated chloride channels (GABACls) are widely expressed in distinct regions of the nervous system of insects (14). Both GABACls (15) and GluCls (16) have variants that occur due to splicing and RNA editing. For the resistant to dieldrin (RDL) GABACls, 4 variants results from alternative splicing at exon 3 and 6. The RDL diversity affects the half-maximal concentration EC50 of GABA (17). In contrast, Bombyx GluCl creates variants by splicing at the exon 3 and 9 sites, and the exon 3 splicing-induced amino acid change affects the receptor density in the membrane (18). To determine the okaramines targets, we isolated cDNAs of the exon 3b/3d variants of Bombyx RDL (BmRDL) and exon 3b/9 (full length) variants of Bombyx GluCl (BmGluCl), since they were expressed most abundantly in the brain and the third thoracic ganglion of the larvae (13). Okaramine B activated only GluCl when tested on the exon 3b/3d variant of BmRDL and exon 3b/9 (full length) variant of BmGluCl expressed in Xenopus oocytes (Figure 3) (13). The BmGluCl-activating potential of okaramines were correlated with both the chloride current-inducing potential in the neurons (r2 = 0.964) (13). Similarly, a strong correlation was observed between the in vivo and in vitro activities (Insecticidal activity vs chloride current inducing activity, r2 = 0.936; insecticidal activity vs GluCl channel-opening activity, r2 = 0.914), suggesting that okaramines selectively activate BmGluCl and thereby induce toxicity in the larvae. Okaramine B was inactive for the human α1β2γ2 GABACl and α1β glycine-gated chloride channel, which suggested its selectivity for insects (13).

Figure 3. Okaramine selectively activates Bombyx GluCl expressed in Xenopus oocytes. Okaramine had no effect on the membrane currents in oocytes expressing the BmRDL, while inducing inward currents in a concentration dependent manner in oocytes expressing the BmGluCl. Applications of GABA and L-glutamate are indicated by horizondal lines, while those of okaramine B are indicated by dashed horizontal lines. Adapted with permission from ref. (13). Copyright 2014 Nature Publishing Group. 128

GluCls are the primary target of the macrocyclic compound ivermectin, which persistently activates the channel in a similar manner to okaramines (19). Hence, we investigated okaramine’s ability to displace [3H]ivermectin at exon 3c/9 (full length) variant expressed in HEK293 cells. Okaramine B reduced [3H]ivermectin binding to the GluCls in a non-competitive manner (20). Thus, the okaramines appear to interact with a different site than ivermectin in the GluCls. However, further studies are required to confirm this mechanism of action by using different GluCls as well as testing with radio-labeled okaramines. Furthermore, it is important to examine, not only the effects of mutations that reduce ivermectin sensitivity of GluCl on the action of okaramines, but also coapplication effects of the two compounds on the ivermectin- and the okaramine-induced response of the GluCls, for understanding the mode of action of okaramines.

Figure 4. Chrodriman B selectively blocks Bombyx RDL GABACl expressed in the silkworm larval neurons. The whole-cell patch-clamp electrophysiology was employed to record the membrane currents in the larval neurons. After recording the response to 30 μM GABA, 1 μM chrodrimanin B was applied for 1 min and then co-applied with 30 μM GABA. Applications of GABA are indicated by horizondal lines, while application of okaramine B is indicated by a dashed horizontal line. Adapted with permission from ref. (21). Copyright 2011 PLoS.

Chrodrimanins: Meroterpenoids Targeting GABACls Chrodrimanins are meroterpenoid compounds that are produced in okara as insecticides by YO-2 of the Talaromyces sp (1). The whole-cell patch-clamp electrophysiology was employed to show that chrodrimanin B had no effect on the membrane current in the silkworm larval neuron when applied alone, but the coapplication of 1 μM chrodrimanin B with 30 μM GABA completely blocked the GABA-induced currents (Figure 4) (21). Furthermore, it blocked the response to 30 μM of BmRDL with an IC50 value of 1.13 nM. The order of the IC50 values of 129

chrodrimanin A (143 nM), B and D (6.01 nM) for BmRDL were in accordance with the order of LD50 values of >100, 10 and 20 μg/g diet, respectively, for the silkworm larvae. Chrodrimanin B also blocked the GABA-induced response of human α1β2γ2 GABACl with an IC50 of 1.5 μM, which was approximately 1,000-fold higher than BmRDL (21).

Concluding Remark Insect-active metabolites of various fungi are being continuously isolated from okara, a soybean-derived medium. Surprisingly, most metabolites have high selectivity for insect ligand-gated ion channels. The discovery of “ifungi”, or induced fungi formed by plant stimulants, may be a potential chemical library for next-generation pesticides.

Acknowledgments Kazuhiko Matsuda (KM) and Makoto Ihara (MI) was supported by KAKENHI (KM, grant number: 17H01472; MI, grant number: 16K21507) from the Japan Society for the Promotion of Science.

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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 Bloomquist, J., 111 Calkins, T., 91 Cens, T., 75 Chahine, M., 75 Charnet, P., 75 Clark, J., 53 Coats, J., ix Collet, C., 75 Furutani, S., 125 Gross, A., ix, 111 Hayashi, H., 125 Ihara, M., 125 Islam, R., 111 Kai, K., 125

Lansky, D., 53 Lorsbach, B., 1 Matsuda, K., 125 Menard, C., 75 Murenzi, E., 53 Ozoe, Y., ix, 19 Piermarini, P., 91 Rousset, M., 75 Soderlund, D., 37 Sparks, T., 1 Symington, S., 53 Thibaud, J., 75 Toltin, A., 53

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Subject Index A Agrochemical discovery, 1 current insecticide research, 5 insecticide market, changes in the percentages, 6f insecticides based on IRAC classification, global sales, 7f early insecticide research, 3 classes of insecticides, timing of the introduction and size, 4f new active ingredients, changes, 3f synthetic organic insecticides, different insecticidal modes, 5f introduction, 2 insect control and delivery systems, options, 2f present and future, approaches to insecticide discovery, 8 approaches to insecticide discovery, 8f competitor inspired (CI) starting points, 9 insecticide discovery, approaches employed, 10 summary and conclusion, 11 insecticidal products, fundamental approaches, 13 recent insecticides, structures, 12f

G Glutamate receptor-cation channel complex, 111 discussion, 121 materials and methods chemicals, 112 glutamatergic agonists, cyclothiazide, and racetams, chemical structures, 113f insects, 114 statistics, 115 toxicity and paralysis bioassays, 114 results adult female Ae. aegypti, intrathoracic injection (LD50), 119t adult mosquitoes, 116 Ae. aegypti, correlation, 119f cyclothiazide and racetam compounds, paralytic activity, 117t

feeding bioassays and Aedes aegypti, 118 glutamatergic agonists, paralytic activity, 116t glutamatergic compounds, intrathoracic injection and oral toxicity, 118t glutamatergic compounds, paralytic activity, 120t glutamatergic compounds with piracetam, synergism, 120 headless larvae paralysis, 115

H Honeybee voltage-gated Ca2+ channels, 75 alternative insecticides target, CaV channels, 85 Ca2+ currents, physiological recordings, 77 voltage-gated Ca2+ channels, in vivo and in vitro characterization, 78f cloning and sequence analysis, 79 Apis mellifera DSC1/CaV4, pore sequence and expression, 82f voltage-gated Ca2+ channels, molecular structure, 81f heterologous systems, functional properties, 83 CaV3 and CaV4, biophysical properties, 84f

I Induced fungi, metabolites introduction, 125 asperparaline A on ACh-induced currents, selective blocking action, 127f fungi in okara, insect-active metabolites produced, 126f meroterpenoids targeting GABACls, 129 okaramines, resolving the enigma, 127 Bombyx GluCl, okaramine selectively activates, 128f silkworm larval neurons, 129f silkworm larval neurons, establishing a patch-clamp recording technique, 127

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Insect control, voltage-gated sodium channels future sodium channel-directed insecticides, 46 discovering new binding sites, 48 natural and synthetic isobutylamide insecticides, structures, 47f targeting unexploited binding sites, 47 past and present, sodium channel-directed insecticides chemical class and target site, insecticide use, 46f DDT, 41 early pyrazoline-based insecticides, structures, 44f indoxacarb, structures, 44f pyrethrins, structures and nomenclature, 40f pyrethroids, 42 pyrethrum, 39 resmethrin, permethrin, deltamethrin and fenvalerate, structures, 42f s-bioallethrin, 41f SCI insecticides, conceptual model of the state-dependent action, 45f SCI insecticides, proposed common core structure, 45f sodium channel inhibitors, 43 U.S. dollar value, pyrethroid use, 43f sodium channels, coevolutionary exploitation, 38 neurotoxin binding sites, 39t voltage-gated sodium channels, 37

M Mosquito gap junctions, 91 dipteran innexins, physiological roles, 97 embryonic development, 98 excretion, 100 immune responses, 99 malpighian tubule of an adult female Ae. aegypti, Inx3 immunolabeling, 101f nervous system function, 98 ovary of Ae. aegypti, Inx3 immunolabeling, 100f reproduction, 99 introduction connexins and innexins, general comparison, 93

dipteran innexins, molecular and functional properties, 95 dual voltage clamp, illustration, 97f gap junctions, brief history, 92 gap junctions, structure, 94f innexin proteins, phylogenetic tree, 96f mosquito borne disease, control, 92 mosquito control, gap junctions as molecular targets, 101 gap junction inhibitors, toxicity, 102f

N Neurolemma-injected Xenopus oocytes, 53 initial neurolemma-injected oocyte assay expected perfomance, confirmation, 58 neurolemma-injected oocytes, TTX-sensitive current, 59f pyrethroids, effects of increasing concentrations, 60f pyrethroids, summary of the effects, 61t rat brain neurolemma tissue, insecticide induced TTX-sensitive inward currents, 57f TTX-sensitive inward current, 56 neurolemma microtransplantation assay concentration-dependent response curves (CDRCs) for pyrethroids, mixed-model regression analysis, 67f data analysis improvements, 66 effects of pyrethroids, summary, 65t incorporation scalar factors, adjusting TTX-sensitive AUC, 64s individual incorporation scalar factor (IISF), 65f variation, identifying the sources, 61 version 2, 62 version 3, 63 Xenopus oocytes, summary of the effects of pyrethroids, 68t neurolemma microtransplantation assay, utility, 68 neurolemma-injected oocyte assay, steps to reduce variability, 69s variation estimates of the logEC50, summary, 70t rat brain tissue, microtransplantation, 54 experimental procedure, overview, 55f

140

P

new-generation GABA receptor NCAs, 26 noncompetitive GABA receptor antagonists, 20f potency of PS-14, dependence, 24f predicted interaction between 3B-4-PIOL and the amino acid, 28f PS-14 inhibition of GABA- and glutamate-induced currents, dose-response curves, 22f TM2 regions of fruit fly, amino acid sequences, 25f phenolamine GPCRs amitraz, 30 octopamine elicited calcium and cAMP signals, 29 octopamine/tyramine GPCR subtypes, 28 second messengers, EC50 values, 30t

Pest control chemicals, 19 GABA- and glutamate-gated chloride channels bicyclophosphates, 21 BPB1 and broflanilide, chemical structures, 27f early-stage GABA receptor research, 20 EBOB, docking, 23f fluralaner and fluxametamide, chemical structures, 26f GABA and competitive antagonists, chemical structures, 27f GABA receptors, molecular pharmacology, 22 GABA receptors, orthosteric ligands, 27 GABA receptors versus inhibitory glutamate receptors, 25

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