Ligand-Gated Ion Channels: Putative Target Sites for Anthelmintic Therapy in Muscle and Intestine Cells of Parasitic Nematodes

Iowa State University Capstones, Theses and Graduate Theses and Dissertations Dissertations

2021

Ligand-gated ion channels: Putative target sites for anthelmintic therapy in muscle and intestine cells of parasitic nematodes

Mark Andrew McHugh Iowa State University

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Recommended Citation McHugh, Mark Andrew, "Ligand-gated ion channels: Putative target sites for anthelmintic therapy in muscle and intestine cells of parasitic nematodes" (2021). Graduate Theses and Dissertations. 18556. https://lib.dr.iastate.edu/etd/18556

This Dissertation is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Graduate Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. Ligand-gated ion channels: Putative target sites for anthelmintic therapy in muscle and intestine cells of parasitic nematodes

Mark McHugh

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Genetics and Genomics

Program of Study Committee: Richard J. Martin, Co-major Professor Jo Anne Powell-Coffman, Co-major Professor Alan P. Robertson Michael J. Kimber Heather M.W. Greenlee

The student author, whose presentation of the scholarship herein was approved by the program of study committee, is solely responsible for the content of this dissertation. The Graduate College will ensure this dissertation is globally accessible and will not permit alterations after a degree is conferred.

Iowa State University

Ames, Iowa

2021

DEDICATION

This dissertation research is dedicated to my loving and caring parents, Norman W.

McHugh and Hazel E. McHugh, to my devoted and loving wife, Deverley S. Scott-McHugh, and my dear siblings Paul N. McHugh and Christene G. McHugh.

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TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS ...... vii

ABSTRACT ...... x

CHAPTER 1. GENERAL INTRODUCTION ...... 1 1.1 Literature Review ...... 3 Parasitic Nematodes ...... 3 Ascaris spp ...... 4 Oesophagostomum spp...... 7 Brugia malayi ...... 9 Nematode Neuromuscular System ...... 11 Muscular system ...... 12 Nervous System ...... 13 Nematode Intestinal Epithelium ...... 16 Anatomy of Ascaris suum Intestine...... 16 The nematode intestine as a therapeutic target for anthelmintics ...... 20 Nicotinic acetylcholine receptors ...... 23 Large N-Terminal Extracellular Domain ...... 24 Transmembrane Domains...... 26 Cytoplasmic Domain and the C-terminus ...... 26 Vertebrate nAChRs ...... 26 C. elegans nAChR ...... 28 Parasitic Nematodes nAChR ...... 32 Ascaris suum...... 32 Oesophagostomum dentatum ...... 35 Haemonchus contortus ...... 37 Brugia malayi ...... 40 Synthesis, assembly, and trafficking of mammalian nAChR ...... 41 Synthesis, assembly, and trafficking of nematode nAChR ...... 43 Anthelmintics and anthelmintic resistance ...... 46 Anthelmintics ...... 46 Mode of Action ...... 47 Imidazothiazoles ...... 47 Tetrahyropyrimidines ...... 49 Macrocyclic lactones (MLs) ...... 52 Benzimidazole (BZs) ...... 54 Anthelmintic resistance ...... 55 1.6 References ...... 58

CHAPTER 2. CHOLINERGIC RECEPTORS ON INTESTINE CELLS OF ASCARIS SUUM AND ACTIVATION OF NACHRS BY LEVAMISOLE ...... 79 2.1 Abstract ...... 79 iv

Keywords...... 80 2.2 Introduction ...... 80 2.3 Materials and Methods ...... 83 2.3.1 Collection and maintenance of A. suum worms ...... 83 2.3.2 Histological preparation of A. suum ...... 83 2.3.3 Intestinal tissue and muscle bag preparation ...... 84 2.3.4 A. suum cDNA synthesis and RT-PCR detection of nAChR subunits ...... 84 2.3.5 Analysis of mRNA levels by Quantitative Real-time PCR ...... 85 2.3.6 Preparation of Formalin-Fixed Paraffin-Embedded (FFPE) tissue for RNAscope ISH ...... 86 2.3.7 RNAscope in situ hybridization Assay ...... 87 2.3.8 Subcellular quantification of mRNA punctate dot analysis ...... 87 2.3.9 Intestinal Calcium Imaging ...... 88 2.4 Results ...... 91 2.4.1 Transverse section of Ascaris suum ...... 91 2.4.2 nAChR subunit mRNA expression in A. suum muscle and intestinal cells ...... 91 2.4.3 Differential expression of nAChR subunits in muscle bags and intestinal tissue ..... 92 2.4.4 RNAscope reveals heterogeneous subcellular distribution of intestinal nAChR subunit mRNAs ...... 92 2.4.5 Subcellular distribution of intestinal nAChR subunit mRNAs ...... 93 2.4.6 Acetylcholine and levamisole stimulate calcium signals in Ascaris intestine ...... 94 2.5 Discussion ...... 97 2.5.1 GAR-1s, nAChRs and a paracrine function ...... 97 2.5.2 Different distributions of nAChRs subunit message in the basolateral and central regions of the columnar intestine cells ...... 98 2.5.3 Possible function of the nAChR subtypes located in the basolateral region of the intestinal cells ...... 99 2.5.4 Possible function of the nAChR subtypes located in the central region of the intestinal cells ...... 99 2.5.5 Levamisole and Cry5B ...... 100 2.6 Conclusion ...... 101 2.7 Declaration of competing interests ...... 101 2.8 Acknowledgments ...... 101 2.9 Author Contribution ...... 101 2.10 Figures and Tables ...... 102 2.11 References ...... 113

CHAPTER 3. ALLOSTERIC MODULATION OF OESOPHAGOSTOMUM DENTATUM NICOTINIC ACETYLCHOLINE RECEPTORS BY MACROCYCLIC LACTONES ...... 119 3.1 Abstract ...... 119 3.2 Introduction ...... 120 3.3 Materials and Methods ...... 123 3.3.1 Molecular Biology ...... 123 3.3.2 Heterologous expression of Ode-levamisole receptors and Ode-pyrantel receptors in Xenopus laevis oocytes ...... 123 3.3.4 Two-microelectrode voltage camp (TEVC) electrophysiology ...... 124 3.3.5 Drugs ...... 125 v

3.3.6 Drug application ...... 125 3.3.7 Data analysis...... 125 3.4 Results ...... 126 3.4.1 Effects of abamectin on Ode levamisole receptors ...... 126 3.4.2 Effects of ivermectin on Ode levamisole receptors ...... 127 3.4.3 Effects of moxidectin on Ode levamisole receptors ...... 127 3.4.4 Effects of abamectin on Ode pyrantel/tribendimidine receptors ...... 128 3.4.5 Effects of ivermectin on Ode pyrantel/tribendimidine receptors ...... 129 3.4.6 Effects of moxidectin on Ode pyrantel/tribendimidine receptors ...... 129 3.5 Discussion ...... 130 3.5.1 Diversity of parasitic nematodes nAChR subtypes ...... 130 3.5.2 The avermectins (abamectin and ivermectin) and the milbeymycin, moxidectin are positive allosteric modulators (PAMs) of the Ode levamisole receptor ...... 131 3.5.3 The avermectins (abamectin and ivermectin) are negative allosteric modulators (NAMs) of the Ode-Pyr/Tbd receptor, while moxidectin exhibits positive allosteric modulatory effects (PAM)...... 131 3.5.4 Use of drug combinations in combating anthelmintic resistance in parasitic nematodes ...... 132 3.6 Conclusion ...... 133 3.7 Declaration of competing interest ...... 134 3.8 Acknowledgement ...... 134 3.9 Author Contribution ...... 134 3.10 Figures and Tables ...... 135 3.11 References ...... 145

CHAPTER 4. ANTHELMINTIC RESISTANCE AND HOMEOSTATIC PLASTICITY: BRUGIA MALAYI ...... 149 4.1 Abstract ...... 149 4.2 Introduction ...... 150 4.3 Methods ...... 152 4.3.1 Parasite maintenance ...... 152 4.3.2 Drugs ...... 153 4.3.3 Dissection ...... 153 4.3.4 Whole-cell recording ...... 153 4.3.5 Calcium imaging and Fluo-3 injections ...... 154 4.3.6 RNA extraction and cDNA synthesis ...... 155 4.3.7 Synthesis and delivery of dsRNA ...... 155 4.3.8 Analysis of Transcript levels ...... 156 4.3.9 B. malayi in vitro Motility Studies ...... 156 4.3.10 Data analysis...... 158 4.4 Results ...... 159 4.4.1 Levamisole produces spastic paralysis, then a flaccid paralysis followed by recovery of motility ...... 159 4.4.2 Calcium fluorescence and effect of levamisole ...... 160 4.4.3 The duration of paralysis and recovery, depend on the cholinergic anthelmintic ... 161 4.4.4 100 µM levamisole inhibits/desensitizes all receptor subtypes ...... 162 4.4.5 unc-38 is upregulated, and nra-2 is downregulated, in levamisole habituated vi

Brugia ...... 163 4.4.6 nra-2 knockdown speeds but unc-38 knockdown slows levamisole desensitization/recovery ...... 163 4.4.7 nra-2 knockdown reduces levamisole currents but not ACh currents ...... 164 4.4.8 Reversal of levamisole habituation and nra-2 expression levels after removing levamisole ...... 164 4.4.9 Recovery and Knockdown of AChR receptor genes...... 165 4.5 Discussion ...... 166 4.5.1 Dynamic and Plastic AChRs in Brugia muscle ...... 166 4.5.2 Desensitization and flaccid paralysis ...... 168 4.5.3 Calcium homeostasis ...... 169 4.5.4 Gene regulation and Recovery ...... 170 4.5.5 Therapeutic significance ...... 170 4.6 Conclusion ...... 171 4.7 Acknowledgements ...... 171 4.8 Author contributions ...... 172 4.9 Competing interests ...... 172 4.10 Figure and Table Legends ...... 172 4.11 References ...... 184

CHAPTER 5. GENERAL CONCLUSION ...... 191 5.1 Future directions ...... 192 vii

ACKNOWLEDGEMENTS

I would like to express my deepest gratitude first and foremost to the Lord Jesus Christ for the wisdom, spiritual guidance, and strength that he has endowed me with to complete my graduate work. Without him, this would be totally impossible. To my major professors Dr.

Richard J. Martin and Dr. Jo Anne Powell-Coffman, words are inadequate to express how grateful I am for agreeing to accept me in their labs when research funds were scarce. I am thankful to them for their expertise, guidance, patience, unfeigned confidence in me and the countless hours they have invested. Through their enthusiasm and ability to approach compelling research problems, my aptitude for research has grown immensely.

Special thanks to my program of study committee (POSC) members: Dr. Alan P.

Robertson, Dr. Heather M.W. Greenlee, and Dr. Michael J. Kimber for agreeing to be on my

POSC and for their support, advice, and guidance during my doctoral studies. In addition, I would like to highlight Dr. Alan Robertson for his altruistic nature, his devotion and passion to solving research problems and his willingness to assist me with refining experimental ideas or thoughts when needed.

I would also like to acknowledge the former coordinator of the Interdepartmental

Genetics Program, Linda Wild (retired) and the staff members the Department of Genetics,

Development and Cell Biology, Danise Jones, Carla Harris, Diane Jespen along with staff members of the Department of Biomedical Sciences, Kim Adams, Seth Shatto and Audra

Hartwigsen for making my tenure comfortable and successful at Iowa State University.

I would like to thank, the College of Liberal Arts and Sciences for their financial support and providing me the opportunity to teach the Biology 211 Lab throughout my doctoral studies.

Sincere thanks to the NIH and the E. A. Benbrook Foundation for Pathology and Parasitology for viii providing funding to my research. Special thanks to my Biology 211 L TA family, Dr. Jim

Colbert, Linda Westgate, Christopher Myers, Jacob Eeling and Dr. Lori Beiderman for their confidence in me and providing a professional and friendly work environment to teach.

I would like to thank my lab mates Dr. Melanie Abongwa, Dr. Shivani Choudhary, Dr.

Sudhanva Kashyap, Dr. Saurabh Verma and Dr. Paul Williams who have been a source of support and assisted me when I needed advice on research matters. To Dr. Ronique Beckford,

Dr. Kwame Matthews, Raye McHugh Jr, Randolph Edmondson, Deniel Simms, Tashine

Bonfield, Kevin Johnson, Sherieka Fearon, Rshana Shurriah and Kishi Anderson, special thanks for their support, encouragement and friendship. A special thank you also to Hyacinth Johnson, who has been a true friend and guardian from an early age until this present day.

To my wife Deverley, I am grateful to her for her prayers, unconditional love, emotional support, encouragement, understanding and patience with me when I arrive home late at nights from lab. To my beloved father Norman McHugh, and mother Hazel McHugh, my brother Paul and late sister Christene thank you for your unfailing love, inspiration, emotional and financial support through all my years of school. I could not have asked for a better family. To Pamella

McKenzie, my adopted sister, thank you for all the love and support that you have displayed to me and my parents.

To my mother-in-law Viviene Powell and her family, my aunt-in-law Jennifer Powell and her family, my uncle-in-law Trevor Powell, sincere thanks to them for their words of encouragement, love, prayers and support. To my uncle, Leon Bertram and his family, I would like to thank them for their prayers, love, support and positive influence, specifically when I just arrived in the US to pursue my studies. Thanks also to my dear aunt, Jean Henry and her family for their constant prayers, love, words of encouragement and support throughout the years. ix

To my church family in White Hall, Jamaica, I would like to thank them for their prayers, unwavering support and their investment in me from my formative years until I became an adult.

To my New Life Church family in Ames, specifically Pastor Daniel Flemming, his wife Amanda

Flemming, his children, Bowen and McKenzie Flemming and his parents, Donald and Darlene

Flemming, I would like to thank them for their prayers, words of encouragement and their love to me. I also want to offer my appreciation to Don and Lynette Hartman, Mark and Ella Tonjum who have provided a family unit for me during my doctoral studies. x

ABSTRACT

The prevalence of parasitic nematode infections are a major human and animal health concern. There are still no effective vaccines available, hence anthelmintic drugs have remained the cornerstone for prophylaxis and treatment. The repertoire of available anthelmintics is limited, with treatment relying heavily on three major chemical classes of anthelmintics. These are the imidazothiazoles/tetrahydropyrimidines (levamisole, pyrantel, morantel, oxantel); benzimidazoles (mebendazole, flubendazole, thiabendazole, albendazole); and the macrocyclic lactones (ivermectin, moxidectin, abamectin), all of which act on parasitic nematode ion channels, except for the benzimidazoles. Ion-channels are crucial components of excitable tissues and valuable targets for anthelmintics.

Prolong treatment and incorrect use of these anthelmintics however, have led to the development of resistance worldwide. Additionally, the development of new anthelmintics is slow-paced, with only three drug classes being developed and approved for animal use in the since the year 2000. This includes the amino-acetonitrile (monepantel), cyclooctadepsipeptide

(emodepside) and the spiroindole (derquantel). Hence, there is an urgent need for the development of new, more effective anthelmintic drugs that can alleviate the morbidity and mortality caused by existing parasite infections. Additionally, significant gaps in our understanding of anthelmintic resistance need to be improved so that we can provide practical solutions on improving drug efficacy and delaying the onset of resistance.

We have confirmed the expression of four nicotinic acetylcholine receptor (nAChR) subunits: Asu-unc-38, Asu-unc-29, Asu-unc-63 and Asu-acr-8 that constitute the putative levamisole receptor in adult female Ascaris suum intestine. We then validated these findings by using RNAscope in situ hybridization to localize the subcellular distribution of the subunits in xi the intestine. Quantitative real-time PCR (qPCR) was also used to confirm the mRNA expression levels of each subunit in both muscle and intestine cells. To determine whether these subunits formed functional receptors that were responsive to cholinergic agonists, we employed calcium imaging. Our calcium imaging results demonstrated that both acetylcholine and levamisole elicited intracellular calcium responses in the intestinal tissue. These findings suggest that the presence of functional nAChRs in the intestine may not be limited to neuromuscular transmission, but an acetylcholine paracrine function. Hence, A. suum intestine can be a suitable target for therapeutic exploitation.

Secondly, we expressed two receptor subtypes, namely Ode-levamisole and Ode pyrantel/tribendimidine from the pig parasite Oesophagostomum dentatum in Xenopus laevis oocytes. We demonstrated that compounds from the two macrocyclic lactone sub-family, the avermectins (abamectin and ivermectin) and the milbemycin, moxidectin are positive allosteric modulators (PAMs) on the Ode levamisole receptor. In contrast, abamectin and ivermectin acted as negative allosteric modulators (NAMs) on the Ode pyrantel/tribendimidine receptor subtype, while moxidectin maintained its PAM action. These findings suggest that the macrocyclic lactones are allosteric modulators of nAChRs and structural differences between each drug or the presence or absence of a subunit, namely ACR-8 may influence the allosteric modulatory effects.

Hence combination therapy that includes macrocyclic lactones and cholinergic anthelmintics, might improve drug efficacy and delay anthelmintic resistance.

Finally, we investigated the adaptation of Brugia malayi to levamisole exposure. We showed that B. malayi recovered motility with loss of sensitivity to levamisole within 4 hours of exposure. Molecular analysis also revealed up-regulation of mRNA levels for one AChR subunit, unc-38 and down-regulation of a gene that encodes for an ER retention protein, nra-2. Patch- xii clamp experiments on 4 hour recovered worms also showed that muscle responses to levamisole had desensitized. Knock down of nra-2 by RNAi resulted in faster recovery in motility, significant reduction in levamisole currents and no change in acetylcholine currents. This suggest that loss of NRA-2 facilitates the insertion of pentameric AChR subtypes in the muscle that are insensitive to levamisole, thus leading to faster recovery in motility in the presence of levamisole. Additionally, simultaneous knockdown of AChR subunits, namely, unc-38, acr-26 and acr-16, inhibited recovery of motility in the worms. These findings are notable and highlights the dynamic mechanisms used to by the parasite to vary AChR subunit composition that generates various receptor subtypes, thus facilitating recovery of motility and insensitivity to anthelmintic exposure (levamisole). This process of habituation can be interpreted as a mechanism of resistance that can be used by parasitic nematodes. 1

CHAPTER 1. GENERAL INTRODUCTION

Parasitic nematodes are among the most prevalent and widespread infectious agents that adversely undermine human and animal health worldwide. Infections are common in tropical and subtropical regions, and disproportionately affect impoverished populations, inflicting crippling morbidity and mortality (Bethony et al., 2006). Furthermore, food security and sustainability of agricultural economics are also impacted negatively (Weaver et al., 2010). In the absence of vaccines and proper sanitation, control strategies rely almost exclusively on the use of a limited number of synthetic anthelmintic drug classes (Fewtrell et al., 2005; Bethony et al., 2006; Hotez et al., 2009; Bartram and Cairncross, 2010). These drugs are generally classified or grouped based on their target site, with the majority acting on the parasite’s nervous system, specifically ion channels. This includes: the macrocyclic lactones, which target glutamate gated chloride channels (GluCls) found only in invertebrates (Cully et al., 1994; Wolstenholme and Rogers,

2005); benzimidazoles, which bind selectively to β-tubulin and inhibit microtubule formation

(Lacey, 1990); imidazothiazole/tetrahydropyrimidines which act as agonists on nicotinic acetylcholine receptors (nAChRs) (Aubry et al., 1970; Aceves et al., 1970; Martin and

Robertson, 2010; Abongwa et al., 2017); cyclooctadepsipeptides, which target latrophilin receptors and SLO-1 potassium channels (Willson et al., 2004; Guest et al., 2007; Holden-Dye et al., 2007; Krucken et al., 2012; Martin et al., 2012); aminoacetonitrile derivatives (AADs) which act as an agonist or antagonist of nAChRs (Kaminsky et al., 2008; Baur et al., 2015; and the spiroindole which act as an antagonist of several subtypes of nAChRs (Robertson et al., 2002;

Woods et al., 2012). Ion channels are suitable drug targets due to their essential role in the normal physiology of the parasite. Inhibition of these channels produce paralysis which inhibits locomotion, feeding and egg production. However, repeated use of these drugs has resulted in 2 the increasing selection of drug-resistant nematode populations, thus leading to the emergence of anthelmintic resistance globally (Prichard, 1994; Sangster et al., 2018; Geerts and Gryseels,

2000). Despite the problem of resistance, the development of novel compounds has been very slow, with the development of only three compounds since the year 2000, one of which has already reported resistance (Nixon et al., 2020). Hence, there is an urgent need to combat the problem of anthelmintic resistance. This can be achieved by improving our understanding of general parasite biology, elucidating drug resistance mechanisms and identifying and characterizing existing and unknown drug targets. Additionally, exploration of non-excitable tissues such as the nematode intestine for ion channels and membrane transporters can be crucial in crippling vital functions that are necessary for the parasite’s survival. Therefore, these PhD studies seek to highlight putative target sites that can be exploited for anthelmintic therapy with the aim of circumventing anthelmintic resistance. The first aim was to confirm the presence of nAChR subunits that constitute the putative levamisole receptor in the intestine (a non-excitable tissue) of the pig parasite, Ascaris suum, and to determine its sensitivity to levamisole. A. suum serves as an excellent model for the human parasite Ascaris lumbricoides and the discovery of this receptor would reveal a putative site of action for anthelmintic therapy. The second aim was to determine the allosteric modulatory effects of the avermectins, namely abamectin and ivermectin and the milbemycin, moxidectin on expressed heteromeric nAChR receptor subtypes from Oesophagostomum dentatum. This was done to improve our understanding on pharmacological mechanisms involved in the improvement of drug efficacy via combination therapy. The third and final aim was to investigate mechanisms of desensitization used by Brugia malayi to adapt to high concentrations of levamisole over an extended period.

This would provide us with a mechanistic explanation for levamisole habituation which can facilitate the emergence of anthelmintic resistance.

1.1 Literature Review

Parasitic Nematodes

Parasitic nematodes have existed for thousands of years. They appeared officially in the first written records from a period of Egyptian medicine, 3000 to 400 BC, particularly in the

Ebers papyrus of 1500 BC discovered at Thebes (Bryan, 1930; Cox, 2002). Documentation of parasitic infections were also made by several civilizations throughout history such as the

Chinese, Greek, Indian, Roman and Arab. Although these civilizations are now a shadow of the past, parasitic nematodes have successfully survived the rise and fall of empires by use of sophisticated mechanisms that gave them an evolutionary advantage.

Parasitic nematodes belong to the phylum Nematoda, where approximately 25,000 species of nematodes are known. As parasites, they spend part of their life cycle in the environment and another aspect in living organisms (host). Here they compete with the host for nutrients, which is essential for their growth and survival. Today, parasitic nematodes are among the most common and widespread infectious agents that negatively impact human and animal health worldwide. They are generally endoparasites that inhabit the host gastrointestinal tract or may be blood borne.

These organisms are prevalent in tropical and subtropical regions of the world, specifically low income and impoverished areas that lack proper sanitation and limited healthcare (Fewtrell et al., 2005; Bartram and Cairncross, 2010; Weaver et al., 2010; Strunz et al., 2014; WHO, 2017). Some of the most common parasitic nematodes include round worms

(Ascaris lumbricoides, Ascaris suum, and Strongyloides stercoralis), whipworms (Trichuris 4 trichiura), hook worms (Ancylostoma duodenale, Necator americanus), filarial worms

(Wuchereria bancrofti, Brugia malayi) and pin worms (Enterobius vermicularis). In our study the parasitic nematodes Ascaris suum, Oesophagostomum dentatum and Brugia malayi are used as model organisms for human parasites. Here we discuss their life cycle, symptoms of infection and diagnosis and treatment of disease.

Ascaris spp

Ascaris spp. are roundworm gastrointestinal parasites that belong to the family

Ascarididae. Major species of Ascaris include: Ascaris lumbricoides which infects humans,

Ascaris suum which infects pigs and Parascaris equorum which infects horses. Both Ascaris lumbricoides and Ascaris suum are very closely related genetically, making Ascaris suum a suitable model nematode in studying the human parasite Ascaris lumbricoides. Ascaris lumbricoides is known to cause Ascariasis, a neglected tropical disease, which infects approximately 800 – 1.2 billion people worldwide and causes 60,000 deaths annually, mainly in children (Pullan et al., 2014; Hadush and Pal, 2016; de Lima Corvinoy and Bhimji, 2018).

Infections are common in southern regions of India, south-east Asia, south and south-west China,

Central and South America and Sub-Saharan Africa (de Silva et al., 2003). The animal species of

Ascaris have also created severe production and economic losses.

Ascaris spp. are large nematodes, with adult females being larger with lengths ranging from 20 – 35 cm, while males are smaller having an average length of 15 – 30 cm. Adult females are also recognized by their characteristic large ovaries which occupy approximately 60% of the worm. Additionally, the posterior end of the female is straight, while that of the male curves ventrally.

The life cycle of Ascaris lumbricoides is monoxenous (direct) with humans as the definitive hosts (Figure 1.1). It commences with female worms laying eggs in the lumen of the 5 small intestine of the host. Adult females generally produce an average of 200,000 eggs per day, which may be fertilized or unfertilized. These eggs are then passed in feces to the environment.

Within the soil, fertilized eggs embryonate and become infective after 18 – 20 days providing that environmental conditions are optimum (low humidity, moist and warm) and insanitary conditions are maintained. In temperate regions, eggs stay dormant in winter and resume development when the temperature is warm and conducive in the spring.

Transmission occurs through the fecal-oral route, where contaminated soil or food containing Ascaris spp. eggs are ingested by another individual. Upon ingestion of the eggs, the eggs are transmitted to the small intestine of the host where first-stage larvae hatch (L1), then molt into second-stage larvae (L2). The larvae then penetrate the intestinal mucosa, enter the bloodstream, and are transported to the liver. From the liver they then migrate to the lungs where they further mature in a period of 10 – 14 days and subsequently migrate from the pulmonary capillaries into the alveoli. Third-stage larvae (L3s) are then coughed up and swallowed, where they enter the small intestine to molt into fourth-stage larvae (L4s) and subsequently develop into adult worms. Adult female worms generally produce eggs approximately 2 – 3 months after infection. Additionally, if the infection is left untreated, adult worms can survive in the host for

12 to 18 months where numerous eggs are continuously produced and passed in feces.

A. lumbricoides infections are generally asymptomatic in individuals who have low to moderate infection. However, larval migration through pulmonary tissues can cause type-1 hypersensitivity reactions, resulting in eosinophilic pneumonia (Loeffler syndrome) associated with fever, cough and wheezing typically 10 – 14 days after infection. Severe infections with adult parasites can manifest symptoms such as abdominal distension and discomfort, intestinal blockage, anemia, malnutrition, diarrhea, and weight loss. Infection in children may lead to 6 impaired growth, impaired cognition, and reduction in school performance due to infrequent attendance.

Source: Accessed on 04-03-21 at 2:42AM CST, from https://www.cdc.gov/dpdx/ascariasis/modules/Ascariasis_LifeCycle_lg.jpg

Figure 1.1 Life cycle of Ascaris lumbricoides

The standard approach for diagnosing ascariasis, is by microscopic identification of A. lumbricoides eggs in stool samples. Treatment is conducted by oral administration of the 7 anthelmintics, namely, albendazole (400mg) and mebendazole (500 mg) in a single oral dose.

Mebendazole may also be administered at 100 mg twice daily for three days in cases where patients manifest uncomplicated infections and are over 12 months old. Alternative treatment may involve the use of ivermectin in a single dose of 150 – 200 µg/Kg.

Oesophagostomum spp.

Oesophagostomum spp. are roundworm parasitic nematodes that belong to the family

Strongylidae. They are the causative agent of oesophagostomiasis, which is characterized by nodule formation in the large intestine of its infected hosts. Hence, members of this genus are classified as nodular worms. Infections in livestock have been reported in several countries worldwide, namely, Brazil, China, Malaysia, West Africa, Indonesia, the Philippines and northern parts of Ghana and Togo. However, high incidence of infection in humans have been reportedly mainly in the northern parts of Togo and Ghana (Storey et al., 2000).

There are several species of Oesophagostomum which may be host specific (infecting one host), whereas others may infect multiple hosts.

The different types of Oesophagostomum species that infect livestock are O. multifoliatum and

O. columbianum (sheep and goats), O. radiatum (sheep, goats and cattle), O. dentatum and O. quadrispinulatum (pigs). Human cases of infection have also been reported where O. bificurum

(primarily a monkey parasite) have been implicated as the causative agent.

All Oesophagostomum spp. have a direct life cycle with no intermediate host involved

(Figure 1.2). Adult worms are approximately 7 – 15 mm long. The life cycle begins with adult females laying eggs in the large intestine of the host, which are passed in feces. The eggs then hatch and release first-stage larvae (L1) at favorable environmental conditions. Female worms lay approximately 5000 eggs per day, similar to other nematodes within Strongylidae (Krepel and Polderman, 1992). First-stage larvae (L1) feed on bacteria in the environment and develop 8 into second-stage larvae (L2) within 24 hours after hatching. The second-stage larvae subsequently molt to the infective third-stage larvae (L3). Infection begins with ingestion of contaminated grass, soil, water or food containing third-stage larvae (L3s). Ingested third-stage larvae (L3) then penetrate the mucosa of the cecum and colon, molt to fourth-stage larvae (L4) and re-enter the lumen of the large intestine where they develop into adults and lay eggs.

Source: Accessed on 04-03-21 at 2:49 AM CST, from https://en.wikipedia.org/wiki/Oesophagostomum#/media/File:Life_cyc.jpg

Fig 1.2 Life Cycle of Oesophagostomum spp.

Infections are generally asymptomatic and may be masked by other signs of parasitic infections, namely ascariasis (Roepstorff and Nansen, 1994; Helwigh et al., 1999). However, severe infections may present clinical signs such as pain in lower right abdomen, protruding abdominal masses, diarrhea, fever, loss of appetite and decreased weight gain (Orihel & Ash 9

1995; Storey et al., 2000). Diagnosis includes postmortem examination, where parasite nodules may be visible in and on the large intestine, along with the presence of the parasites in the lumen.

Additionally, microscopic examination of fecal samples may be conducted to identify parasite eggs.

Treatment in animals is achieved by use of broad spectrum anthelmintics such as levamisole, benzimidazoles and several macrocyclic lactones that are effective at eradicating adults and larval stages. Narrow spectrum anthelmintics may also be used namely, pyrantel, morantel, closantel. However, these drugs are mainly effective against adults and not larval stages. In cases of human infections, therapy typically involve administering a single 400 mg dose of albendazole or pyrantel pamoate for adult patients, while children are given 200 mg. For severe cases where abscesses or fistulae arise from Dapaong tumors, drainage is generally performed along with antibiotic and albendazole treatment.

Brugia malayi

Brugia malayi, a clade III filarial nematode, is one of the three causative agents of lymphatic filariasis (elephantiasis) in humans. Lymphatic filariasis is an important neglected tropical disease (NTD) that has infected 120 million individuals and poses a serious threat to over 893 million people in 49 countries worldwide (WHO, 2021). Additionally, it has disfigured and incapacitated 40 million individuals (WHO, 2021). The disease is restricted to rural areas of

South and Southeast Asia.

Brugia malayi adult worms are threadlike in appearance with females having a length of 8 cm while males are 2 cm long (Strickland, 1991). They possess only longitudinal muscles and move in an S-shape motion (Decraemer et al., 2003). The life cycle of this parasite begins with adult worms producing millions of microfilariae in lymphatic vessels of the infected host, which circulates in the blood (Figure 1.3). Adult worms can live in human lymphatic vessels for up to 6 10 to 8 years. Mosquitoes of the genera Culex, Anopheles and Aedes which serve as vectors, become infected during blood feeding from an infected individual, leading to the ingestion of microfilariae. The microfilariae then penetrate the gut wall of the mosquito and migrate to the muscles of the thorax where they undergo maturation into the infective larval stage over a period of 7 – 21 days. Infective larvae then migrate to the proboscis of the mosquito and await transfer to a suitable host. Transmission to other humans then occur when an infected mosquito bites an individual, resulting in the deposition of mature larvae on the punctured human skin, which then enters the body. The larvae then migrate to the lymphatic vessels where they develop into adult worms and continue the cycle.

Infections are generally asymptomatic, but damages may still be caused to the lymphatic and immune system. Chronic symptoms can lead to lymphatic vessel blockage, thus resulting in fluid accumulation and massive swelling of the infected tissues (scrotum, limb, breasts etc.) known as elephantiasis. This disfigurement or deformity is accompanied by social stigma, inability to work, loss of income, depression and poverty. Diagnosis involves microscopic identification of microfilariae in blood smear taken at night due to the nocturnal periodicity of microfilariae. Additionally, serological techniques may also be employed by use of the rapid immunochromatographic IgG4 cassette test that measures anti-filarial IgG4 levels in the blood of an infected individual. Treatment of elephantiasis is dependent on the prevalence of other diseases such as onchocerciasis (river blindness) and loiasis (Loa loa). In regions where onchocerciasis is prevalent, ivermectin (200 mcg/kg) and albendazole (400 mg) are administered together, whereas diethylcarbamazine citrate (DEC) (6 mg/kg and albendazole (400 mg) is given in the absence of onchocerciasis. Areas that have loiasis, albendazole (400 mg) is administered alone twice annually. 11

Vector control is also conducted to reduce the spread of the parasite through use of insecticide- treated nets, residual indoor spraying along with other measures.

Source: Accessed on 04-03-21 at 3:03 AM CST, from https://www.cdc.gov/dpdx/lymphaticfilariasis/modules/B_malayi_LifeCycle_lg.jpg

Figure 1.3 Life Cycle of Brugia malayi

Nematode Neuromuscular System

The neuromuscular system of Ascaris spp. has been extensively studied by light and electron microscopy. Like all nematodes, the neuromuscular system is peculiar in the sense that 12 elongate extensions project from the muscle cells to the nerves, whereas in vertebrates, nerves send processes that extend to muscle cells (Stretton, 1976). Overall, the neuromuscular system is crucial to the survival of parasite, thus providing them with the movement required for maintaining their position in the host intestine. Here we briefly highlight the neuromuscular system of the human and pig parasite, Ascaris lumbricoides and Ascaris suum respectively, which are very similar anatomically.

Muscular system

The somatic muscle of A. suum is among the earliest studied and consists entirely of longitudinally oriented cells. Each body wall muscle cell is divided by two lateral lines into ventral and dorsal halves and composed of three morphologically distinct and spatially separate components (Rosenbluth, 1965; Stretton, 1976) (Figure 1.4). This includes the spindle fiber, the belly and the arm (Figure 1.5). The muscle spindle or fiber contains an obliquely striated contractile apparatus that forms a long tube attached to the hypodermis (Rosenbluth, 1965; del

Castillo et al., 1989). The belly or bag region is a turgid balloon-like structure that is continuous with the core of the striated spindle fiber (Rosenbluth, 1965). It measures 200 µm in diameter and lies in the pseudocoelomic space. It serves as an endoskeleton and contains the nucleus.

Additionally, it stores glycogen particles which can become depleted during starvation of the nematode (Rosenbluth, 1965). Next is the muscle arm which is a thin process that originates from the base of the muscle bag and extend towards the syncytium which lies over the nerve cord. Upon arrival of the muscle arms at the syncytium, the end of the arms divides into several finer processes called fingers (del Castillo et al., 1989). Additionally, tight junctions are also formed between adjacent fingers in the syncytium, thus providing electrical coupling between adjacent cells (DeBell et al., 1963; Rosenbluth, 1965; del Castillo et al., 1967).

Neuromuscular junctions are also formed between the fingers of the arms which forms the syncytium and the nerve cord.

Nervous System

The nervous system of A. suum commences in the head region where there is a nerve ring that surrounds the pharynx. Arising from the nerve ring are two major nerve cords, the dorsal and ventral nerve cords. Anatomical studies have shown that the ventral nerve cord consists of cell bodies of motorneurons (Stretton, 1978). These motorneurons are separated into seven anatomical types based on the distribution of their dendrites and axon. This includes dorsal excitatory DE1, DE2 and DE3; ventral excitatory VE1 and VE2; dorsal inhibitory DI and ventral inhibitory VI (Stretton, 1978). Connections are also made between the ventral and dorsal nerve cords via commissures, namely, one left hand commissure and three paired right-hand commissures (Figure 1.6) (Stretton, 1978; Martin, 1993).

Source: Martin 1993, p. 18.

Figure 1.4 (A) Diagram of A. suum worm showing the anterior region of the transverse section of B (B) Transverse section of A. suum showing the structures that constitute the somatic muscle. 14

The pattern of the commissures repeat itself along the length of the body of the worm, with each repeat forming five segments and each segment containing eleven motoneurons. These eleven motorneurons include: DI, DE2 and DE3 which occurs once in each segment and DE1, VI, V1 and V2 occurring twice in each segment (Stretton et al., 1978).

Source: Del Castillo et al., 1967, p. 264.

Figure 1.5 Illustration showing a cross-section of the muscle cell of Ascaris spp. Here we have the muscle bags (m.b.), muscle arms (m.a.), tight junction (t.j.), neuromuscular junction (n.m.j), nerve cord (n.c), muscle fiber (m.f.), syncytium (syn), hypodermis (hyp) and cuticle (cut). 15

In addition to the major nerve cords, there are also minor lateral and sublateral nerve cords in the anterior region of the worm. This includes: the left and right ventral sub-lateral cords and the left and right dorsal sub-lateral cords (Stretton, 1976). These minor nerve cords are critical for complicated head movements of the worm which is required for feeding. The tail region also consists of four minor nerve cords which are the left and right ventral nerve cords and left and right dorsal nerve cords. Together these nerve cords are vital for complex tail movements during copulation.

Source: Martin, 1993, p. 15.

Fig 1.6 Diagram of the anatomical organization of Ascaris suum dorsal and ventral nerve cord in one segment. Each segment of the ventral nerve cord has six interneurons and eleven motorneurons along its length. The seven anatomical types of motorneurons are shown as DI: dorsal inhibitory, VI: ventral inhibitory, DEI: dorsal excitatory 1, DE2: dorsal excitatory 2, DE3: dorsal excitatory 3, VI and V2: ventral excitatory. There are also left and right commissures which are axons of motorneurones that are single or paired and pass round the body of the nematode. 16

Nematode Intestinal Epithelium

The intestinal epithelium is one of the major organs of Ascaris suum. It is complex structurally and conducts critical functions that are vital to the parasite’s survival. Additionally, it is emerging as a valuable research model that can be exploited for anthelmintic therapy. We use this section of the literature review to give concise information on the anatomy of A. suum intestine and highlight its significance as a target site for current and prospective anthelmintic drugs.

Anatomy of Ascaris suum Intestine

The intestine of Ascaris suum consist of a single layer of columnar epithelial cells that enclose the lumen (Sheffield, 1964) making it suitable for absorption and digestion of nutrients.

It is divided into three regions, namely, the anterior, middle and posterior intestine (Harpur,

1977) (Figure 1.7). The anterior intestine starts immediately posterior to the pharynx with a constriction known as the pharyngeointestinal junction. This is then followed by a thin-walled expansion chamber that is held in place by delicate attachments that consists of connective tissue and the body wall muscle (Wright et al., 1972; Harpur, 1977). At the end of the anterior intestine is a coarctation or narrowing, which marks the junction of the anterior and mid intestine (Harpur,

1977). This is generally proximal to the genital pore of the female worm. The characteristic feature of the mid intestine is that it lies freely amongst the reproductive organs and lacks connective tissue attachments and compression by body wall muscle cells. Next is the posterior intestine which lacks body wall muscle attachments at the mid-intestine and posterior intestine junction (Wright et al., 1972; Harpur, 1977). However, it has continuous attachment of body wall muscle at its posterior region. Termination of the intestine occurs shortly at the junction with the rectum. 17

Source: Hapur, 1977, p. 1116.

Figure 1.7 Schematic diagram of a dissected female Ascaris suum nematode. The intestine is depicted as a long tube laying in the center of the nematode (black). It starts posterior to the pharynx and ends at the anus. Labels on the left show the three subdivisions and nomenclature proposed by the authors for the intestine. Labels on the right indicate other anatomical organs of the nematode.

The internal anatomy of the intestinal epithelium starts in the lumen. Cells which are ventral or dorsal to the lumen in the anterior region are approximately 50µm in height and 10µm in width (Sheffield, 1964). In contrast, cells at the lateral edges of the lumen are generally shorter

(Sheffield, 1964). At the apical membrane, are slender, finger-like projections known as the 18 microvilli (Sheffield, 1964) (Figure 1.8). This structure was initially considered to be a cuticular border (Bilek, 1909), a layer composed of rows of vacuoles (Quack, 1913) or cilia (Bilek, 1909;

Hetherington 1923; Mueller, 1929) in different species of Ascaris.

Source: Kessel et al., 1961, p. 115.

Figure 1.8 Schematic diagram showing the ultramicroscopic view of the intestinal epithelium of Ascaris spp. The structures are as following: microvilli (MV), rootlets (R), terminal bars (desmosomes) (TB), terminal web (FN), cell membrane infoldings (IM), endoplasmic reticulum (ER), mitochondria (M), lipid inclusions (L), nucleus (N), granules (G), infoldings of the basal region of the plasma membrane (I), basal lamella (BL) and mesentrial membrane (PM).

However, electron microscope studies of the intestine at higher resolution revealed the structure to be protoplasmic projections in the form of microvilli (Bretschneider, 1950; Chitwood &

Chitwood, 1950; Bretschneider, 1954; Beams & Anderson, 1957). Each microvillus is approximately 6 to 7µm long and 0.08µm in diameter (Kessel et al., 1961; Sheffield, 1964).

Inside each microvillus are longitudinal fibers or meshwork of filaments that extend into the superficial apical cytoplasm of the cell forming rootlets. The microvilli also have a filamentous coat of a mucopolysaccharide called glycocalyx (Maggenti, 2012). This provides the microvilli a large surface area for absorption and digestion. Kessel et al. (1961) estimated that the microvilli increased the surface by 75 to 90 times in A. suum.

Beneath the microvilli is the terminal web which has numerous fibrous networks perforated by cytoplasmic connections to the remainder of the cell cytoplasm. Its width spans approximately 3µm wide from one terminal bar (desmosomes) to the next (Kessel et al., 1961).

The terminal bars attach adjacent intestinal cells and are observed close to the lumen surface of the intestine. Additionally, the plasma membrane of each cell interdigitates with adjacent neighboring cells. These interdigitations are fold-like and are visible beneath the terminal web and at the most basal region of the cell. The spaces within the interdigitations provide the opportunity for widening, thus facilitating larger intercellular spaces when necessary (Kessel,

1961). They are also believed to be critical for strengthening adjacent cell-cell connections

(Fawcett, 1955).

Beneath the terminal web is the cytoplasm which provides a habitat for several subcellular organelles. This includes: the mitochondria, endoplasmic reticulum, ribosomes, Golgi apparatus, lipid droplets, glycogen granules and lamellar bodies. The mitochondria are numerous and densely concentrated below the terminal web, in the apical region of the cell, and are fewer 20 in numbers at the basal region. They vary in shape, where some form dense elongated beaded strands, while others are oval, depending on the physiological processes at the time of fixation

(Sheffield, 1964). The endoplasmic reticulum is randomly distributed throughout the cell and are in close association with large deposits of glycogen. The lamellar bodies are believed to be critical in the destruction and elimination of waste or involved in the synthesis of glycocalyx that coats the microvilli.

At the basal position of the intestine is the subspherical to ovoid nucleus. This is followed by the basolateral membrane (basal lamella) which separates the intestine from the pseudocoelomic space (body cavity). The basal membrane is approximately 7µm in width and has a collagen-like composition. Its inner layer is relatively dense and thick, while its outer layer is less dense and thin (Kessel, 1961). The plasma membrane of the basal region is highly folded in a manner similar to the nephron tubules of the crayfish (Anderson & Beams, 1956) or the kidney tubules of vertebrates (Pease, 1955).

The nematode intestine as a therapeutic target for anthelmintics

The nematode intestine is involved in several important functions which includes secretion of enzymes, absorption of digested nutrients, innate immunity, protection against xenobiotics and ion transport. The single cell structural layer and strategic location of the intestine, posterior to the pharynx and central in the body cavity of nematodes, provides increased accessibility of drugs that could be efficiently transported to various tissues in the nematode. Additionally, impairment of this essential organ by anthelmintic therapy would be detrimental to the nematode’s survival.

Several studies have shown the effects of synthetic anthelmintics, such as the benzimidazoles on the intestine of multiple parasitic nematode species. Van Den Bossche and De

Nollin (1973) conducted biochemical studies which revealed that mebendazole (MBZ) inhibited 21 the uptake and transport of low molecular nutrients (carbohydrates and amino acids) in the intestine of A. suum. The authors observed that inhibition of glucose by mebendazole subsequently resulted in a significant reduction in the uptake of amino acids such as proline, glycine, methionine, and triggered depletion of glycogen content in the muscle cells of the nematode (Van den Bossche & De Nollin, 1973). These findings were considered significant, since A. suum has been previously proven to rely entirely on carbohydrate catabolism for their energy supply in anerobic environments (Saz, 1970). In another study, in vivo treatment of A. suum with a low concentration of mebendazole, revealed irreversible morphological alterations in subcellular organelles of the intestine (Borgers & De Nollin, 1975). These degenerative modifications included: swelling and disruption of microvilli, reduction in apical secretion granules and glycogen content, and excessive accumulation of autophagic vacuoles throughout the cytoplasm (Borgers & De Nollin, 1975). These results confirm that impairment of the intestinal epithelium can lead to the interruption of normal biochemical processes in the parasite, thus deeming the intestine as an important target for anthelmintic therapy.

Jasmer et al. (2000) also conducted in vivo studies that showed that the intestine of the parasitic nematode, Haemonchus contortus, was hypersensitive to fenbendazole treatment (FBZ).

This resulted in disintegration of the anterior intestine, DNA fragmentation in the intestinal nuclei in an apoptosis-like process and dispersal of secretory vesicles throughout the cytoplasm of the intestine of the worms. The authors also postulated that the mechanism of efficacy of fenbendazole on the intestine was due to inhibition of secretory vesicle transport, via depolymerization of microtubules. Taken together, these results show that anthelmintic therapy is not limited to the inhibition of neuromuscular function but can induce effects on other tissue types, namely the intestine, thus impairing its function and leading to the parasite’s demise. 22

In addition to commercially available anthelmintics, natural plant extracts have also shown anthelmintic properties against the nematode intestine. Williams et al. (2014) has shown that fourth-stage A. suum larvae (L4) treated with condensed tannins were motionless and experienced direct structural damage to the cuticle and intestine. This was confirmed by transmission electron microscopy of the intestine which showed that the microvilli in the brush border was significantly destroyed, accompanied by the formation of vacuoles in the intestinal tissue. However, the mechanism by which this was achieved is still unknown.

Bacterial toxins namely the crystal (Cry) proteins, have also been implicated in exhibiting significant anthelmintic actions on the intestinal tract of nematodes. One such Cry protein,

Cry5B, is a pore-forming peptide that is synthesized by the gram-positive soil bacterium,

Bacillus thuringiensis (Bt). Cry5B’s mode of action involves activation by intestinal protease in the nematode. This activated form of Cry5B then binds to a nematode specific cadherin (CDH-8) which leads to internalization and formation of pores, thus damaging and compromising the integrity of the intestinal epithelium (Peng et al., 2018). Wei et al. (2003) demonstrated that exposure of the free-living nematode, C. elegans to Cry5B resulted in toxicity. Briefly, worms were subjected to intoxication, developmental and gut morphology assay. It was observed that nematodes fed E. coli that expressed Cry5B, experienced intoxication which was manifested as inhibition of growth, lack of pharyngeal pumping, lethargy and death. In addition, intestine morphology revealed significant damage where there was shrinkage of the intestine from the body wall muscle and vacuole formation. These results provide confirmation that Cry5B exhibit anthelmintic properties that directly target the intestine of the nematode, similar to that of insect pests. Cry5B have also been deemed effective against nematodes that are resistant to nAChR agonists. Hu et al. (2010) used developmental inhibition and mortality assays to show that C. 23 elegans mutants that were resistant to levamisole, pyrantel and tribendimidine were, hypersusceptible to Cry5B. Additionally, the authors also reported that Cry5B combination with tribendimidine or levamisole, showed strong synergy, thus increasing therapeutic efficacy.

Cry5B’s nematocidal activity is not limited to free-living nematodes, but also extends to parasitic nematodes. Capello et al. (2006) reported that hamsters infected with the human hookworm, A. ceylanicum had an 89% reduction of parasite burden when treated with Cry5B. In another study, Urban et al. (2010) demonstrated that in vitro and in vivo exposure of A. suum fourth-stage larvae (L4) and adults to Cry5B caused intoxication. The authors also showed that

A. suum expressed at least one glycolipid receptor which have been proven to be a target receptor for Cry5B. Finally, the effect of recombinant Cry5B (rCry5B) was investigated on multiple life stages (L1, L3, free-living adult stage and parasitic female stage) of the human parasitic threadworm Strongyloides stercoralis in vitro (Charuchaibovorn et al., 2019). Results showed that feeding worms with recombinant Cry5B led to significant susceptibility by reduction in motility, growth impairment and death of a high percentage of the worms (Charuchaibovorn et al., 2019). Taken together, these results confirm that the nematode intestine is a valuable target for commercially available anthelmintic drugs, natural plant extracts and the bacterial toxin,

Cry5B. This provides an avenue for new drugs that may be developed to target this site of action.

Nicotinic acetylcholine receptors

Nicotinic acetylcholine receptors (nAChRs) are members of the Cys-loop ligand gated ion channels (LGICs) super-family, which includes glycine, 5-hydroxytryptamine (5-HT3) and

γ-aminobutyric acid (GABA) receptors (Lester et al., 2004; Millar, 2006). Additionally, this superfamily also includes histamine-gated chloride channels (HisCl), glutamate gated chloride channels (GluCl) and serotonin-gated chloride channels, all of which are found only in invertebrates. The nAChRs are critically involved in both mediating and modulating synaptic 24 transmission in the nervous system of vertebrates and invertebrates, making them vital targets for therapeutic drugs.

The most detailed structural information on nAChRs was first derived from electron microscopy studies of the electric organ of the Torpedo californica (Changeaux et al., 1996).

These experiments facilitated the creation of a refined 4 Å model of the receptor which was then entirely resolved at 4.6 Å (Unwin, 2005). This revealed the shapes and dimensions of the molecule, the ligand binding sites and ion channel organization. From this we know that nAChRs are pentameric complexes in which five homologous subunits co-assemble to form a central cation pore like the staves of a barrel (Nys et al., 2013; Corringer et al., 2012, Sine & Engel,

2006), (Figure 1.9A and B). Additionally, there are four types of subunits, namely, α, β, γ and δ.

All protein subunits exhibit basic homologies throughout their sequences and have lengths of approximately 500 amino acids. Each subunit is also structurally similar, with a common architecture that consists of a large N-terminal extracellular domain, a transmembrane domain, a cytoplasmic or intracellular domain, and a short extracellular carboxyl domain (Zoli et al., 2018)

(Figure 1.9C).

Large N-Terminal Extracellular Domain

The N-terminal domain has N-glycosylation sites that are present throughout each subunit. Several loop regions are also present which are vital to receptor function. This includes six loops (A – F) and the Cys-loop (two cysteine residues separated by 13 amino acids).

Furthermore, classification of subunits as either α or non-α is based on specific features shared in loop C, where α subunits contains two vicinal cysteine residues at positions 192 and 193. In contrast, non-α subunits lack this feature. Loops A – F have also been implicated in forming the ligand (acetylcholine) binding site (Brejc et al., 2001). This has been proven by mutagenesis and affinity labelling studies which showed that two separate parts of the extracellular domain are 25 involved in acetylcholine (ACh) binding. This includes both the “principal” (positive side) and

“complementary” (negative side components). The principal component consists of loops A, B and C of one subunit, while the complementary component is formed by loops D, E and F of the adjacent subunit (Brejc et al., 2001; Zoli et al., 2018). The vicinal cysteine residues in loop C are also essential for acetylcholine binding. Collectively, these loops form the binding site for acetylcholine which facilitates the transient opening of the channel.

Source: Zoli et al., 2018, p. 339.

Figure 1.9 Diagram of the structure of the nicotinic acetylcholine receptor. A) Structure showing the side view of the pentameric receptor with the putative ACh-binding sites and the influx and outflow of ions upon channel activation. B) Aerial view of the five subunits arranged around a central pore, with transmembrane 2 (M2) lining the channel (dark grey). C) Image of a single nAChR subunit embedded in the membrane. The large N-terminal extracellular domain is followed by the four transmembrane domains (M1 – M4), a large cytoplasmic loop between M3 – M4 and the short extracellular C-terminal domain.

Transmembrane Domains

Each nAChR subunit consists of four α-helical transmembrane domains (TM1 – TM4).

TM1 is located between the ACh binding site of the extracellular domain and TM2 (Figure

1.9C). TM1 is also considered to be essential in acting as a linkage between ligand binding and channel gating based on its location. TM2 of all five subunits are involved in forming the lining of the pore of the ion channel which contains the gate and determinants for selectivity (Figure

1.9B). There are three negatively charged rings which includes an extracellular ring, a narrow middle ring that selectively filters the channel and a cytoplasmic ring on the intracellular region

(Martin et al., 1997). All three rings are lined with negative amino acid residues, thus allowing the conduction of cations through the channels while repelling anions and large cationic structures. TM3 is located immediately before the cytoplasmic (intracellular) domain, while

TM4 is located at the C-terminal end of the cytoplasmic domain. Collectively, TM1, TM3 and

TM4 are crucial in coiling around each other and forming an outer ring that shields the inner ring of the TM2 domains from the membrane lipids (Figure 1.9B).

Cytoplasmic Domain and the C-terminus

The cytoplasmic loop is located between TM3 and TM4 (Figure 1.9C). Its sequence is highly variable and contains phosphorylation sites that are involved in the modulation of nAChR function and receptor localization (Figure 1.9C) (Zoli et al., 2018). The C-terminus is short and begins immediately after the TM4 region and are located on the extracellular surface.

Vertebrate nAChRs

There are seventeen known nAChR subunits found in vertebrates, namely, α1 – α10, β1 –

β4, γ, δ and ε (Millar & Gotti, 2009). These subunits assemble to form heteromeric or homomeric receptor subtypes that are diverse in stoichiometry and pharmacologically distinct.

Vertebrate nAChRs are also further categorized as muscle nAChRs or neuronal nAChRs based on their location.

Muscle type nAChRs are well characterized due to their simple homogenous system.

They are heteromeric with distinct subunit stoichiometry that are developmentally regulated due to stage specific changes in gene expression pattern of some subunits. The embryonic muscle prior to innervation is comprised of four kinds of subunits, 2α1, β1, γ and δ, while the adult muscle nAChR consists of 2α1, β1, δ, ε (Mishina et al., 1986). Muscle type nAChRs are located postsynaptically at the neuromuscular junction and possess two binding sites for ACh and other cholinergic agonists. This may be formed at interfaces between a α-subunit or adjacent ε/γ or δ subunit, thus providing different binding affinities for agonists.

Neuronal type nAChRs are considerably diverse in comparison to the homogeneous population of muscle type nAChR at the neuromuscular junction. There are twelve subunits (α2

– α10 and β2 – β4) that have been identified which are predominantly expressed in the central and peripheral nervous system. These subunits can form functional homomeric or heteromeric receptors but is strongly dependent on the subunit type and stoichiometry.

Heterologous expression in Xenopus oocytes have revealed that any of the three α subunits (α2, α3, α4) when co-expressed with either β2 or β4 are able to co-assemble to form functional heteromeric receptors. However, they were unable to form functional homomeric receptors when expressed on their own. By contrast, α7, α8 and α9 subunits are a distinct branch of nAChR subunits that can generate functional homomeric channels based on several lines of evidence from electrophysiology experiments. Overall, C. elegans and Drosophila have been used as suitable models that provide evidence of co-assembly and diversity of vertebrate nAChR subunits. 28

C. elegans nAChR

The free-living nematode, Caenorhabditis elegans have been widely used as a model organism for studying nematode biology and a resource for antiparasitic drug development.

Research using this nematode is due to the worm’s completely sequenced genome, simple nervous system, rapid life cycle and ease of generating transgenic strains, sensitivity to the majority of commercially available anthelmintics and the ability to perform forward and reverse genetics.

C. elegans has the largest and most diverse nAChR gene family in a single species, with at least 32 nAChR subunits (Figure 1.10). These 32 subunits are further divided into five groups based on sequence homology, with each group named after the first member to be discovered.

This includes: DEG-3, UNC-29, UNC-38, ACR-8 and ACR-16. A summary of each group and subunit expression pattern can be seen in, Table 1.1. It should also be noted that there are 22 α subunits while 10 are non-α. To further understand the functionality of these subunits, several studies have been conducted via mutagenesis screening, GFP-expression, electrophysiology and pharmacological assays. These experiments yielded information on various functional subtypes of nAChRs in C. elegans, namely, levamisole-sensitive, nicotine-sensitive and the amino- acetonitrile-sensitive (monepantel) receptors.

Levamisole is one of the most widely studied cholinergic anthelmintics. It selectively paralyzes nematodes by activating nAChRs in their body wall muscles, thus producing contraction and spastic paralysis. The road to discovery of the levamisole-sensitive receptor in C. elegans, involved multiple experimental adventures. Initial work by Fleming et al. (1997) provided evidence that heterologous expression of UNC-38, UNC-29 and LEV-1 subunits in

Xenopus oocytes, formed a functional levamisole-sensitive receptor. Moreover, the current amplitude was small, which may not have corresponded to physiological receptors in C. elegans, 29 thus indicating missing subunits. Further work by Richmond and Jorgensen, (1999) revealed that receptor subunit mutations resulted in reduced levamisole-activated currents in C. elegans body wall muscles, which also conferred levamisole resistance.

Subsequent in vivo recording experiments on the body wall muscle of C. elegans further demonstrated that there were two nAChR subtypes, namely a levamisole-sensitive receptor which required both unc-29 and unc-38 subunits and another receptor that was activated by nicotine and acetylcholine (Richmond & Jorgensen, 1999). Notably, rapid desensitization was observed upon nicotine application which is characteristic of α7 receptors. In addition to lev-1, unc-29 and unc-38, the C. elegans unc-63 subunit gene has been shown to confer levamisole sensitivity (Culetto et al., 2004).

Source: Brown et al., 2006, p. 620.

Figure 1.10 Tree display of C. elegans and vertebrate ACh-gated ion channel subunits superfamily. Of interest are the five core groups of the nAChR subunits which are highlighted in 30 blue, green, pink, red and yellow. Subunits that show high sequence homology with known nAChR subunits that do not fall within the core groups are classified as orphans.

Boulin et al. (2008) then revealed that the L-type AChR required five subunits, namely, UNC-

38, UNC-29, UNC-63, LEV-1 and LEV-8 (also called ACR-13) and three ancillary proteins

(UNC-50, UNC-74 and RIC-3) to form a functional receptor in Xenopus oocytes. The ancillary proteins were found to be necessary for assembly, and trafficking of the receptor (Eimer et al.,

2007; Halevi et al., 2002; Haugstetter et al., 2005).

Table 1.1 Summary of the five core groups of nAChR subunits in C. elegans along with their expression pattern. (Source: Holden-Dye et al. 2013, p. 607)

Nicotine-sensitive AChRs (N-AChR) or levamisole-insensitive nAChRs have been found to be homopentameric, unlike the levamisole receptor which is heteropentameric. This has been 31 proven through early work by Ballivet et al. (1996) which showed that Ce21, now ACR-16, formed a functional homomeric receptor in Xenopus oocytes that responded to both nicotine and acetylcholine, but not to levamisole. It was also observed that the receptor was more sensitive to nicotine than acetylcholine, but nicotine acted as a partial agonist of ACR-16, whereas with α7, it is a full agonist. Rapid desensitization was exhibited by the receptor and was blocked by the synthetic compound, dihydro-β-erythroidine (DHβE) and the plant toxin, d-tubocurarine which is similar to that of the α7 receptor. However, N-AChR was insensitive to α-bungarotoxin (BTX) and methyllycaconitine (MLA) which are potent antagonists of the α7 nAChRs. RIC-3 has also been shown to be needed for the functional expression of the receptor in Xenopus oocytes

(Ballivet et al., 2002). These findings were remarkable, but further work was conducted to investigate the subunit composition of the N-AChR receptor. Touroutine et al. (2005) used microarray analysis to confirm that acr-16 was the only essential subunit for this receptor.

Another subunit, acr-8, was also identified, but was found not to be required. Hence, the N-

AChR receptor was deemed as a homopentamer which is similar to the vertebrate homologue α7 which are also homomeric in vivo.

The development of monepantel, an amino-acetonitrile derivative that causes spastic paralysis of body wall and pharyngeal muscles of nematodes, has further highlighted the diversity of the nematode nAChR. Monepantel is known to target key members of the C. elegans

DEG-3 group, namely ACR-20 and ACR-23 which have been shown to form homomeric channels. Rufener et al. (2013) conducted heterologous expression of C. elegans ACR-23 in

Xenopus oocytes which revealed that the receptor was responsive to acetylcholine, choline, nicotine and the anthelmintic monepantel. In addition, choline responses were also strongly enhanced by monepantel. Monepantel, is known to cause spastic paralysis of body wall and 32 pharyngeal muscles. As for ACR-20 it was observed that low concentrations (<1 nM) of monepantel exhibited positive allosteric modulatory effects on the receptor, whereas higher concentrations (> 0.1 µM) produced direct agonist effects on the receptor (Baur et al., 2015).

Overall, these subunits are nematode-specific, thus making them suitable targets for anthelmintics.

Parasitic Nematodes nAChR

Extensive work has been conducted to characterize nAChR receptors in parasitic nematodes. This has shed light on the complexity of these channels to diversify their subunit combination yielding various receptor subtypes that assist in their survival in the mammalian host. Most anthelmintic drugs that are currently in use, are designed to act on nAChR on the somatic muscle of the parasite with the goal of causing paralysis of the worm and quick removal from their predilection site. Some of the major parasites that have been investigated are Ascaris suum, Oesophagostomum dentatum, Haemonchus contortus and Brugia malayi.

Ascaris suum

Ascaris suum (clade III) is a gastrointestinal nematode of pigs and a model organism for the human parasite, Ascaris lumbricoides. The relatively large size of A. suum makes it amendable to dissection of individual tissues and electrophysiology experiments. Numerous studies have been conducted to improve our understanding of the mode of action of anthelmintic drugs, which has allowed different nAChR subtypes to be characterized in A. suum muscle.

Patch-clamp recordings from A. suum muscle have revealed the presence of three different pharmacological subtypes of cholinergic receptors, namely the L-, N- and B-subtype (Robertson et al., 2002; Martin et al., 2003; Martin et al., 2004; Qian et al., 2006). Each subtype is designated based on their relative sensitivities to selective agonists and antagonists, channel conductance and mean open times. 33

Source: Wolstenholme and Neveu, 2017, p. 669.

Figure 1.11 Representation of C. elegans and prominent parasitic nematodes nAChR subtypes based on putative subunit arrangements expressed in Xenopus laevis oocytes. The most potent agonist (blue arrows) or antagonist (red arrows) are indicated below each receptor subtype. When different agonists can activate a receptor subtype, they are indicated from the most potent to the least potent (left to right). Agonists includes levamisole (Lev); pyrantel (Pyr); bephenium (Beph); tribendimidine (Tbd); nicotine (Nic); oxantel (Oxa); and monepantel (Mon). The antagonist is derquantel (Der).

The L-subtype is selective for levamisole and pyrantel, and sensitive to the antagonist paraherquamide; the N-subtype is sensitive to nicotine, oxantel and methyridine, and relatively insensitive to the antagonist paraherquamide; and the B-subtype that is selective for Bephenium and can be selectively antagonized by 2-deoxy-paraherquamide (derquantel). It should be noted however, that levamisole activates all three receptor subtypes at high concentrations (30 µM).

The single channel conductance and mean open times are respectively, 24 pS and 0.6 ms for the

N-type, 35 pS and 0.9 ms for the L-type, and 45 pS and 1.3 ms for the B-type.

Although various nAChR subtypes have been identified and characterized in A. suum muscle, the search for the specific subunits involved in forming functional channels are still ongoing. Progress has been made however with the N-and L-subtype receptors. Subunit components of the N-subtype receptor was revealed through heterologous expression in Xenopus laevis oocytes of a homomeric receptor consisting solely of A. suum ACR-16 subunits and RIC-3 ancillary factor. The results demonstrated that the receptor was sensitive to nicotine, acetylcholine, epibatidine, cytisine and 3-bromocytisine, and had desensitization rates that varied with each agonist. However, the Asu-ACR-16 receptor was insensitive to α-bungarotoxin, unresponsive to α7 positive allosteric modulators and had lower calcium permeability when compared to the α7 receptor. It was also observed that the receptor was not a target to the cholinergic anthelmintic agonists (levamisole, pyrantel, tribendimidine, bephenium, morantel, methyridine and thenium) except for oxantel that evoked a small response. Though this receptor is not the target of currently available cholinergic anthelmintics, slight differences in its pharmacology when compared to the mammalian α7 receptor makes it a suitable target for novel anthelmintic compounds. 35

The quest for determining the subunits that constitute the L-AChR in A. suum was undertaken by Williamson et al. (2009) who identified the colocalization of unc-29 and unc-38 orthologues in the muscle cell membrane of A. suum. These findings suggested that both subunits are components of the L-AChR receptor. Further experiments involving the reconstitution of both subunits in Xenopus oocytes, revealed the formation of functional heteromeric receptor subtypes which responded to levamisole, acetylcholine, and nicotine (Williamson et al., 2009).

However, levamisole and nicotine sensitivity were contingent on the stoichiometric ratios of unc-

38 and unc-29 cRNA injected into each oocyte. Injection of oocytes with unc-38 and unc-29 in a

1:5 ratio, resulted in levamisole acting as a full agonist while nicotine displayed partial agonist response. The receptor was also sensitive to pyrantel but unresponsive to oxantel. In contrast, reversing the ratio (5:1 unc-38: unc-29), resulted in levamisole acting as a partial agonist, nicotine as a full agonist, oxantel evoked response but there was no pyrantel response.

Collectively, this highlights the formation of two distinct receptor subtypes where their pharmacology is directly related to the stoichiometry of the subunits. Hence, the 1:5 (unc-38: unc-29) yielded an L-AChR subtype and the 5:1 (unc-38: unc-29) produced an N-AChR subtype receptor. Unfortunately, further characterization of the receptor was not futile due to lack of reproducibility. Hence, these in vitro experiments may not necessarily reflect the endogenous receptor in A. suum muscle. In this regard it is highly likely that the A. suum L-AChR may consists of additional subunits similar to that of Oesophagostomum dentatum and Haemonchus contortus which require a total of four subunits (3α and 1 non-α) along with three the ancillary proteins (RIC-3, UNC-50 and UNC-74).

Oesophagostomum dentatum

Oesophagostomum dentatum (clade V), a strongylid parasitic nematode of pigs, has been investigated extensively through in vivo and in vitro electrophysiology analysis to gain 36 knowledge of the pharmacology of its nAChR receptors. Early work by Martin et al. (1997) using the patch clamp technique on O. dentatum muscle vesicles, revealed the presence of two main subtypes of levamisole-activated channels, namely, G35 and G45. These L-AChR subtypes were distinguished based on the mean open time and channel conductance, G. Additionally, the presence of other subtypes was also suggested, G25 and G55, thus predicting four subtypes of L-

AChR. Interestingly, the G35 subtype of O. dentatum shares similar channel conductance to that of the L-AChR from A. suum. Subsequent studies by Robertson et al. (1999), expanded on the initial observations of Martin et al. (1997) and demonstrated the presence of four levamisole- sensitive channels on the body wall muscle of O. dentatum, namely, G25, G35, G40 and G45.

Furthermore, comparative analysis of the channel properties in levamisole sensitive (SENS) and resistant (LEVR) isolates showed loss of the G35 channel in levamisole-resistant (LEVR) parasites, which was present in levamisole-sensitive (SENS) parasites (Robertson et al., 1999).

Although the presence of various subtypes of nAChRs have been demonstrated by these single channel recordings, the subunit composition and stoichiometry of the receptor subtypes were still unknown. Heterologous expression of different nAChR subunit combinations yielded four pharmacologically distinct nAChR subtypes with differences in cholinergic anthelmintic sensitivities. These include the pyrantel (Pyr)-nAChR which consists of UNC-63 and UNC-29 subunits; the pyrantel/tribendimidine (Pyr/Tbd)-nAChR composed of UNC-63, UNC-29 and

UNC-38 subunits; the acetylcholine (ACh)-nAChR composed of UNC-63, UNC-29 and ACR-8 subunits; and the levamisole (Lev)-nAChR composed of UNC-63, UNC-29, UNC-38 and ACR-

8 subunits (Buxton et al., 2014). Additionally, the L-type receptor expressed in Xenopus oocytes showed similar conductance and mean open times as the endogenous L-type receptor in vivo.

Taken together, these findings demonstrate the heterogeneity of nAChRs in parasitic nematodes, 37 which is facilitated by variation in stoichiometric arrangements, thus contributing to different sensitivities to various anthelmintics.

Haemonchus contortus

Several studies have been conducted on the trichostrongylus nematode, Haemonchus contortus, to determine the subunit components of its nAChR subtypes. H. contortus, O. dentatum and C. elegans have been shown to be members of the same phylogenetic group (Clade

V), (Mitreva et al., 2004). Advancement in genomic sequence information has provided important information that has enhanced our understanding of shared traits exhibited by these nematodes at the molecular level. C. elegans orthologues of unc-38, unc-29, unc-63 and lev-1 have been identified in H. contortus and other trichostrongylus parasitic nematodes, namely,

Teladorsagia circumcincta and Trichostrongylus columbriformis (Neveu et al., 2010). It was also observed by these authors that there are four copies of unc-29 in H. contortus and other trichostrongylus nematodes, which highlights the diversity of nAChR receptors in parasitic nematodes. In addition, although unc-38 and unc-63 are widely conserved in parasitic nematodes, there has been a lack of lev-8 (a key subunit in C. elegans L-type receptor) conservation in H. contortus and other parasitic nematodes (Williamson et al., 2007; Neveu et al., 2010). However, transcriptomic studies have identified acr-8 in H. contortus and have deemed it important in levamisole resistance (Fauvin et al., 2010). The presence of a truncated isoform of unc-63 (Hco-unc-63a) has been observed in levamisole resistant H. contortus isolates along with the full-length transcript Hco-unc-63b (Neveu et al., 2010). This truncated isoform of unc-63 has also been implicated in levamisole resistance in H. contortus.

With this background information, experiments have been conducted to determine the subunit composition of the levamisole receptor in H. contortus. Heterologous expression in

Xenopus oocytes revealed the formation of two functional levamisole-sensitive nAChR with two 38 different pharmacology, based on the presence or absence of ACR-8 (Boulin et al., 2011). These receptors were classified as Hco-L-AChR1 and Hco-L-AChR2. The Hco-L-AChR1 subtype constituted the H. contortus gene products of UNC-38, UNC-29, UNC-63 and ACR-8 and conserved ancillary factors, RIC-3, UNC-50 and UNC-74. This receptor subtype was most sensitive to levamisole, insensitive to nicotine and dihydro-β-erythroidine (DHβE) and showed weak sensitivity to pyrantel. In contrast, the Hco-L-AChR2 subtype consisted of UNC-38, UNC-

29 and UNC-63 subunits along with the ancillary factors, RIC-3, UNC-50 and UNC-74. This receptor subtype exhibited reduced activation by levamisole and increased sensitivity to pyrantel, nicotine and dihydro-β-erythroidine (DHβE) (Boulin et al., 2011). These findings highlight the fickle nature of parasitic nematodes nAChR, where changes in the subunit composition or stoichiometry yields a diversity of receptor subtypes that is different in pharmacology and sensitivity.

H. contortus has also been known to have other types of functional heteromeric receptors that are not limited to the levamisole-sensitive (Hco-L-AChR1) and pyrantel sensitive receptor

(Hco-L-AChR2). Courtot et al. (2015) has provided evidence of co-expression of ACR-26, a α- subunit and ACR-27, a non-α subunit in the body muscle cells of H. contortus at different developmental stages. Additionally, the authors also co-expressed both subunits (ACR-26/27) along with ancillary factors, RIC-3, UNC-50 and UNC-74 in Xenopus laevis oocytes. This resulted in the reconstitution of a functional receptor that was highly sensitive to morantel and pyrantel, but insensitive to levamisole and nicotine. This was classified as the Morantel-type receptor (M-type). It was also observed that the M-type receptor was formed from ACR-26 and

ACR-27 subunits of Parascaris equorum, a distantly related parasitic nematode of horses.

Interestingly, the pharmacology of the receptor was similar to that of Hco-26/27, where Peq- 39

26/27 was sensitive to morantel and pyrantel but showed an exception for which Peq26/27 was found to be sensitive to levamisole (partial agonist activity).

The diversity of H. contortus nAChR was further investigated with the development of the amino-acetonitrile derivative, monepantel. Moreover, reports of resistance to monepantel by parasitic nematodes led to the discovery of the acr-23 gene in C. elegans and mptl-1, a homologous gene in H. contortus (Kaminsky et al., 2008; Rufener et al., 2010). Further phylogenetic analysis also showed that mptl-1 is more closely related to C. elegans acr-20, than acr-23 (Rufener et al., 2010). C. elegans acr-23 (betaine receptor) and acr-20 are members of the nematode specific DEG-3 family, and form homomeric nAChRs. Heterologous expression of

H. contortus mptl-1 revealed the formation of a functional homomeric channel that was sensitive to betaine and choline. However, monepantel was shown to act as a positive allosteric modulator of betaine and choline currents at low concentrations (<1nM), and a direct agonist that irreversibly opened the channel at higher concentrations (> 0.1 µM), (Baur et al., 2015).

Additionally, the C. elegans acr-20 receptor also experienced irreversible opening by monepantel, but allosteric modulation was decreased when compared to that of the Hco-mptl-1 and C. elegans acr-23 receptors (Baur et al., 2015). Taken together, these studies highlight the pharmacologically complexity on nAChRs, where various receptor subtypes are formed that are diverse in sensitivity to each anthelmintic.

Monepantel is known to act on members of the DEG-3 family of homomeric nAChR, which includes the C. elegans ACR-20 subtype and ACR-23 subtype receptors (Rufener et al.,

2013; Baur et al., 2015). The H. contortus monepantel-sensitive receptor is encoded by Hco- mptl, also a member of the DEG-3 family of receptors. The molecular target of monepantel in 40 other species has not been defined and orthologues of Hco-mptl-1 are not obvious in the genomes of other parasitic nematodes.

Brugia malayi

Unlike the other parasitic nematodes discussed, Brugia malayi, is a filarial nematode that reside in the lymphatic tissues of humans. Studies have shown that B. malayi possess several subtypes of nAChR receptors that are sensitive to various cholinergic agonists (Robertson et al.,

2013; Verma et al., 2017). The success of these findings is attributed to the employment of the patch clamp technique and the Worminator. B. malayi is a member of the Clade III phylogenetic group, similar to A. suum. Robertson et al. (2013) conducted whole-cell patch clamp recordings on adult B. malayi muscle cells which showed responses to acetylcholine, tribendimidine, pyrantel, levamisole and bephenium. These authors further investigated the desensitizing effects of levamisole on the motility of adult female B. malayi. It was observed that worm motility decreased by 85% in 5 mins, complete paralysis was achieved in 10 mins and active motility was restored in 60 mins, similar to that of control worms (Robertson et al., 2013).

Subsequent single cell PCR analysis by Verma et al. (2017) led to the identification of unc-38, unc-29, unc-63, acr-8, acr-16, acr-26 amplicons in the muscle cells of B. malayi. The authors also confirmed expression of unc-27 in whole B. malayi worms but were unable to replicate detection of the subunit in single muscle cells consistently (Verma et al., 2017).

Expression of these subunits reveal the presence of four different putative nAChRs (L-, M-, N- and P) in the muscle cells of B. malayi. In the same study worms were exposed to various cholinergic agonist, namely, morantel, pyrantel, levamisole and nicotine and experienced paralysis upon exposure to each anthelmintic. Additionally, it was also observed that dsRNA knockdown of unc-38 + unc-29 (L- and/or P- type receptors) abolished motility, whereas knock down of acr-16+acr-26 (M- and/or N- type receptors) had little effect on the motility phenotype 41

(Verma et al., 2017). Whole-cell patch clamp recordings on the muscle cells of the parasite illustrated the presence of functional nAChRs where the cells showed sensitivity (inward current response) to morantel, pyrantel, acetylcholine, levamisole, nicotine and bephenium. It was also shown that levamisole, pyrantel and bephenium current responses were inhibited in unc-29+unc-

38 dsRNA-treated worms (Verma et al., 2017). Finally, these authors also showed that monepantel had no effect on B. malayi muscle cells which indicates the absence of monepantel receptors (Verma et al., 2017). Overall, these results collectively underscore the presence of heterogeneous nAChR subtypes in B. malayi muscle, based on the observed sensitivity of the parasite to various cholinergic anthelmintics. Additionally, it also highlights shared similarities with A. suum with the presence of the N-, L-, P- and B-subtype of receptors.

Synthesis, assembly, and trafficking of mammalian nAChR

The nAChRs of the mammalian muscle have been intensely studied, and provides valuable information concerning the synthesis, assembly, and trafficking of nAChRs to the plasma membrane (Colombo et al., 2013). Synthesis and assembly of individual nAChR subunits begins in the lumen of the endoplasmic reticulum (ER). Simultaneously, ER membrane associated ribosomal complexes interact with the nascent subunit polypeptide, where it undergoes covalent modifications (signal peptide cleavage, oxidation of disulfide bonds and N- glycosylation of specific residues), folding and interactions with ER-resident chaperone proteins

(Merlie et al., 1982; Blount & Merlie, 1988; Blount et al., 1990). ER-resident chaperones include

BiP/GRP78 which promotes subunit folding by interacting with the initial segment of the nascent polypeptide that protrudes from the sec61 channel (gated protein translocation channel at the ER)

(Wanamaker & Green, 2007). This interaction is achieved through continuous binding and unbinding cycles, thus inhibiting water and ions from moving through the pore. ERp57 is also another critical ER-resident chaperone protein that interacts with the polypeptide to promote 42 proper subunit folding, along with the ER-membrane protein, calnexin (Wanamaker & Green,

2007). Collectively, these proteins retain immature polypeptides within the ER to ensure proper folding and successful maturation, by stabilizing the polypeptide, thus increasing the availability of subunits for receptor assembly. Additionally, they are also involved in regulating receptor assembly in the ER. Moreover, the criteria that is used in nAChR subunit assembly into functional pentameric structures is not properly understood.

Proteins that are misfolded or unassembled are retained in the ER and ultimately targeted by an ubiquitin monomer that covalently attaches to lysine residues of the protein (Christianson

& Green, 2004). These ubiquitinated proteins are then retro-translocated from the ER into the cytosol for degradation by the proteasome (Christianson & Green, 2004). Another important ER- resident protein that is involved in the maturation and export of nAChRs from the ER is RIC-3

(resistance to inhibitors of Cholinesterase 3). The activity of RIC-3 is dependent on its level of expression in the ER. Alexander et al. (2010) reported that down-regulation of RIC-3 promotes

α7 subunit assembly and subsequent surface delivery, whereas up-regulation of RIC-3 suppresses α7 surface delivery and retains the subunit in the ER. The cytosolic loop of nAChR subunits is also essential in determining proper subunit folding and influencing the level of cell surface receptors (Colombo et al., 2013).

Recruitment and transport of assembled nAChR receptors from the ER to the Golgi is carefully regulated by specific ER retention/retrieval signals that are present in the transmembrane domain and cytosolic loop of each subunit. This ER retention signal is a conserved motif in the transmembrane domain, whereas the retrieval signal in the cytoplasmic loop is a di-basic sequence (Arg313 – Lys314). The assembly of the subunits into pentameric receptors, or the binding of a subunit to a chaperone protein such as 14-3-3, buries the 43 retention/retrieval signal, thus enabling successful export from the ER to the Golgi (Jeanclos,

2001). However, if the subunits are unassembled, the signals are recognized by the COPI protein complex, which retain them in the ER (Keller et al., 2001).

The cytosolic loop is not only involved in receptor folding and negative regulation of receptor trafficking by the presence of a retrieval signal (di-basic signal) (Keller et al., 2001). It also has signals that promote ER export. This entails conserved hydrophobic residues that facilitate interactions with the ER export machinery (Gotti et al., 2005). Recognition and binding of this motif to COPII proteins allows receptor recruitment at the ER exit sites and trafficking to the Golgi in COPII vesicles (Mancias & Goldberg, 2008). Within the Golgi apparatus, further quality control and processing is done, which includes the modification of N-linked oligosaccharides on δ and γ subunits, into more complex forms (Gu et al., 1989; St John, 2009;

Rudell et al., 2020). VILIP-1 a Ca2+- sensor protein then promotes the exit of the mature receptors from the Golgi apparatus to the cell surface membrane (Zhao et al., 2009).

Synthesis, assembly, and trafficking of nematode nAChR

The mechanisms of synaptic expression and function of nematode nAChRs have been of great significance to parasitologists and pharmaceutical companies over the past decades. Of specific interest is the biogenesis and trafficking of nAChRs in nematodes which can be exploited for therapeutic purposes (Figure 1.12). Numerous ground-breaking findings have given us the tools to carry out pharmacological experiments by use of Xenopus oocytes and cell lines to reconstitute and express nAChRs. However, knowledge concerning the involvement of specific proteins and sequential events that occur during nAChR synthesis and trafficking from the ER to the cell surface membrane is still sparse. The levamisole-receptor (L-AChR) has been studied extensively in C. elegans, and provides valuable insights into some of the proteins that are involved in the synthesis, trafficking and expression of functional nAChRs. 44

Nematode nAChRs are synthesized in the ER, similar to that of mammalian nAChRs.

Within the endoplasmic reticulum (ER), a subunit of a conserved ER membrane complex

(EMC), EMC-6, have been implicated in stabilizing L-AChR subunits before or during their assembly (Richard et al., 2013). Additionally, EMC-6 is also considered to be involved in protein folding, to ensure that the subunits have the correct conformation. Proteins that are misfolded, are generally retained in the ER or exported to be degraded. Overall, the EMC complex functions in protein folding, subunit maturation and also provides a translocation pore that enables the insertion of membrane proteins in the ER (Chitwood & Edge, 2019). Another important step in nAChR subunit production is post-translational modification which occurs simultaneously with folding and assembly of subunits. Proteins such as UNC-74 and CRLD-1, are also ER resident proteins that have been suggested to serve as a disulfide isomerase (PDI) where they catalyze disulfide bond formation, promote subunit stability before or during assembly and facilitate subunit maturation. Knockdown of the CRLD-1 homolog (Creld1) in mice resulted in reduced surface expression of mammalian skeletal muscle nAChR

(D’Alessandro et al., 2018), thus emphasizing the significance of this protein in nAChR maturation.

RIC-3, an ER resident chaperone protein, has also been implicated in affecting the stability of newly formed subunits and are involved in assembly, maturation and trafficking of nAChR receptors (Halevi et al., 2002; Castillo et al., 2005; Alexander et al., 2010). Another important pair of proteins that interact with each other are NRA-2 and NRA-4. Both proteins are considered to play a role in selecting subunits during assembly of L-AChRs. Almedom et al.

(2009) reported that a loss-of-function mutation in NRA-2 led to increased synaptic expression of one subunit, ACR-8, while reducing expression of UNC-38. 45

Subunits that have been assembled are then ready to be trafficked from the ER to the Golgi apparatus. Here, UNC-50, a conserved membrane protein that resides in the Golgi apparatus, directs the L-AChRs to the plasma membrane. Studies have shown that in the absence of UNC-

50, L-AChR subunits degrade in a lysosome-dependent manner, thus confirming the significance of this protein in the trafficking and functional expression of L-AChRs (Eimer et al., 2007).

Another important partner that is involved in nAChR receptor trafficking, is the WNT signaling pathway. Jensen et al. (2012) showed that mutations of several components of the WNT signaling pathway led to the accumulation of ACR-16 in the muscle arms of C. elegans. The authors concluded that WNT signaling facilitates the translocation of ACR-16 from intra-cellular pools to the cell surface membrane (Jensen et al., 2012). Additionally, the WNT signaling pathway is also critical in enabling plasticity in the neuromuscular junction (NMJ), thus influencing ACR-16 synaptic expression when there is increased neuronal activity (Jensen et al.,

2012).

Upon arrival of the nAChR receptors at the plasma membrane, there are auxiliary proteins that have been implicated in interacting with the receptors, thus influencing their properties. This is different in contrast to the mammalian system where there are no auxiliary proteins implicated with nAChRs that are successfully expressed at the plasma membrane.

However, in C. elegans, the presence of EAT-18 was proven crucial for EAT-2 (pharyngeal receptor) expression (McKay et al., 2004; Choudhary et al., 2020). MOLO-1 has also been shown to affect L-AChR properties (Boulin et al., 2012). Finally, proper localization of the receptors are pivotal for efficient synaptic transmission. Proteins such as LEV-10, LEV-9, OIG-

4, MADD-4 and EAT-6 have all been shown to co-localize at the NMJs where they form a clustering complex with the L-AChR receptor, thus playing a role in synaptic localization (Gally 46 et al., 2004; Gendrel et al., 2009). RSU-1 protein also functions in eliminating formation of extrasynaptic L-AChR clusters.

Source: Treinin and Jin, 2020, p. 8.

Figure 1.12 Proteins involved in the maturation and functional expression of C. elegans levamisole receptor (L-AChR).

Anthelmintics and anthelmintic resistance

Anthelmintics

Anthelmintic drugs are chemical agents that are developed for use in controlling parasitic helminth infections in human and veterinary medicine. In the absence of vaccines, these drugs have played a critical role to reduce transmission and ease symptoms of infection. They can be administered orally, by injection or as pour-on depending on the organism been treated (Sangster 47 et al., 2018). There are several classes of anthelmintics, namely, imidazothiazoles, tetrahydopyrimidines, benzimidazoles, salicylanilides, macrocyclic lactones, the amino- acetonitrile derivatives, the cyclooctadepsipeptides, and the spiroindoles. Each class of anthelmintics has a unique mode of action, thus allowing them to act on specific target sites within the parasite. Anthelmintics are designed to either kill the parasite, or cause expulsion from the body of the host without causing significant damage to the host (Martin et al., 1997). Here we review the mode of action of four major classes of anthelmintics that have been widely used worldwide and are relevant to the studies undertaken by our group. In addition, updates are also provided on the development of anthelmintic resistance which serves as a global problem.

Mode of Action

Imidazothiazoles

Imidazothiazoles are broad spectrum anthelmintics that are efficacious against nematodes but lack anthelmintic activity against trematodes and cestodes. The first drug that was synthesized and introduced to the market in this class of anthelmintics was tetramisole. It is an aminothiazole derivative that has a racemic mixture of two optical isomers in equal amounts, namely, S (-) tetramisole (L – tetramisole or levamisole) and R (+) tetramisole (D-tetramisole)

(Thienpont et al., 1966; Raeymaekers et al., 1966, 1967), (Figure 1.13). Aceves et al. (1970) discovered the mode of action of this compound which involved selective binding of tetramisole to nicotinic acetylcholine receptors on the nematode muscle cells, thus leading to contraction and spastic muscle paralysis. Later experiments revealed that the L-isomer was more potent than the

D-isomer which consequently led to the development of levamisole from the pure L isomer (Van den Bossche & Janssen, 1967; Thienpont et al., 1969).

The effects of levamisole on parasitic nematodes have been studied thoroughly via electrophysiology and motility assays, thus confirming the mode of action of this class of 48 anthelmintics. Robertson and Martin, (1993) conducted single channel patch clamp experiments on muscle vesicles from A. suum. It was observed that application of levamisole (1 – 90 µM) to the extracellular surface, selectively activated cation channels that had mean open times in the range 0.80 – 0.25 ms and an average channel conductance of 32.9 ± 1.23 pS. Additionally, open channel block and desensitization was observed at higher concentrations (30 µM and 90 µM) of levamisole.

Source: Van den Bossche & Janssen, 1967, p. 1781.

Figure 1.13 Chemical structures of S (-)-tetramisole (levamisole) (A) and R (+)-tetramisole (B)

In another study Qian et al. (2006), observed that levamisole activated a nAChR subtype in A. suum muscle that had a similar mean open time of 0.9 ms and a channel conductance of 35 pS as that previously reported. Patch-clamp work on levamisole sensitive and levamisole resistant O. dentatum also revealed that the sensitive isolate had the G35 subtype, while it was absent in the resistant isolate. Heterologous expression of O. dentatum L-type receptor in

Xenopus laevis oocytes also showed levamisole selectivity with a single channel conductance of

35 pS (Buxton et al., 2014). Blanchard et al. (2018) also reconstituted a L-subtype receptor for

H. contortus in Xenopus oocytes that was levamisole selective.

Motility assays have been used to demonstrate levamisole’s inhibitory effects on parasitic nematode motility. Verma et al. (2017) reported that exposure of adult B. malayi parasites to 10 49

µM levamisole resulted in spastic paralysis that was sustained. This observation was also previously seen in B. malayi as reported by Robertson et al. (2013). However, worms experienced recovery within 60 mins when treated with 30 µM levamisole. This brief period of transient paralysis and recovery was also seen in B. malayi adults and microfilariae, A. suum and

Nippostrongylus brasiliensis worms when exposed to high concentrations of levamisole (Coles,

1975; Robertson et al., 2013; Mostafa et al., 2015). Taken together, these experiments highlight the mode of action of levamisole by targeting nAChRs in several parasitic nematodes.

Additionally, observations of transient paralysis in worms treated with high concentrations of levamisole, shows that paralysis may not be fatal, but adequate time is allowed for quick parasite removal from their site of predilection via host peristalsis.

Tetrahyropyrimidines

Tetrahydropyrimidines are a class of drugs that have similar mode of action as imidazothiazoles agonists (Aubry et al., 1970; Martin, 1997). They act by selectively binding as agonists at synaptic and extrasynaptic nAChRs on muscle cells of nematodes resulting in contraction and spastic paralysis (Robertson & Martin, 1993). Members of this class of drugs include pyrantel, morantel and oxantel (Figure 1.14). Several studies have been undertaken that have confirmed the mode of action of these drugs on parasitic nematode nAChRs.

Pyrantel was the first of the compounds to be introduced in 1966 where its initial use was as a broad-spectrum drug for livestock and companion animals. Single-channel recordings from the muscle vesicles of A. suum revealed that pyrantel activated cation-selective channels which had similar properties to acetylcholine activated channels that was previously studied, thus confirming pyrantel as a nAChR receptor agonist (Pennington & Martin, 1990; Robertson et al.,

1994). The authors also showed that pyrantel acted as an open channel-blocker of nAChRs, similar to levamisole at high concentrations (Robertson et al., 1994). Classical pharmacological 50 experiments on A. suum muscle strips have also revealed the presence of an L-subtype receptor that was selective not only to levamisole but also pyrantel (Martin et al., 2004). Coexpression of nAChR subunits, namely UNC-29 and UNC-63 from O. dentatum in Xenopus oocytes produced a functional receptor that was sensitive to pyrantel (Buxton et al., 2014). In the same study, the authors also included an additional subunit, UNC-38 with a final combination of O. dentatum

UNC-29, UNC-63 and UNC-38 which reconstituted a functional receptor that was sensitive to both pyrantel and tribendimidine, but not levamisole (Buxton et al., 2014). H. contortus has also been shown to have a functional heteromeric receptor (Hco-L-AChR2) that is pyrantel sensitive based on successful reconstitution in Xenopus oocytes. In B. malayi, Verma et al. (2017) conducted motility assays that showed that adult female B. malayi motility was inhibited by pyrantel, leading to spastic paralysis. In addition, the authors also conducted whole-cell patch clamp experiments on B. malayi muscle where sensitivity to pyrantel was observed, thus illustrating the presence of functional P-type channels on B. malayi muscle cells (Verma et al.,

2017).

Source: Martin, 1997, p. 13.

Fig 1.14 Chemical structures of (A) pyrantel, (B) Morantel and (C) Oxantel

Morantel a methyl ester analogue of pyrantel was the second drug that was produced in this class of anthelmintics as a nematocidal compound for veterinary use. Experiments at the single-channel level, have demonstrated that morantel causes the activation and blocking of the 51

L-subtype receptors in A. suum (Evans & Martin, 1996). Other studies have also shown shared sensitivity between morantel and pyrantel on the morantel-type (M-type) receptor. Courtot et al.

(2015) demonstrated that coexpression of H. contortus ACR-26 and ACR-27 in Xenopus oocytes led to the expression of a functional receptor that was highly sensitive to both morantel and pyrantel, but insensitive to oxantel and levamisole. It was also observed in the same study, that the Parascaris equorum ACR-26/27 receptor showed strong sensitivity to both morantel and pyrantel, while levamisole exhibited partial agonist activity (Courtot et al., 2015). Abongwa et al. (2016) have also shown that morantel is a non-competitive antagonist of the A. suum ACR-16 receptor expressed in Xenopus oocytes. Moreover, it was also observed that the manner in which morantel conducted inhibition was through open channel blocking (Abongwa et al., 2016). In B. malayi, morantel has been shown to inhibit worm motility along with eliciting sensitivity in the somatic muscle of the parasite, thus illustrating the presence of an M-type receptor in B. malayi

(Verma et al., 2017).

Oxantel, the most recent drug of this group, is an m-oxyphenol derivative of pyrantel

(McFarland & Howes, 1972). This was developed due to lack of pyrantel efficacy on whipworms. It possesses unique characteristics when compared to the other compounds in this class by not having an exclusive receptor subtype. Martin et al. (2004) reported that the N-type receptor in A. suum, is preferentially activated by oxantel and other compounds. This is different in contrast to pyrantel that was preferential to the L-subtype. However, both oxantel and pyrantel have shared pharmacological activities on A. suum muscle cells where oxantel was observed to cause open channel block, similar to pyrantel (Dale & Martin, 1995). Oxantel has also been shown to be efficacious against one species of hookworm but lack efficacy against other species.

Heterologous expression of A. caninum ACR-16 in Xenopus oocytes showed that the receptor 52 was insensitive to oxantel (Choudhary et al., 2019). In another study A. americanus and A. ceylanicum, ACR-16 receptor expressed in Xenopus oocytes showed that only A. ceylanicum was sensitive to oxantel, while A. americanus ACR-16 lacked sensitivity (Kaji et al., 2020). Hansen et al. (2021) also discovered Tsu-ACR-16 as sensitive molecular target for oxantel in the pig whipworm, Trichuris suis where oxantel acted as a full agonist on the receptor. Pyrantel also activated the receptor moderately, while nicotine elicited minor responses.

Macrocyclic lactones (MLs)

Macrocyclic lactones (MLs) are a potent class of anthelmintics that have been widely used in the control of nematode parasites in humans, livestock and companion animals

(Campbell & Benz, 1984). These compounds are hydrophobic and are derived from soil microorganisms of the genus Streptomyces (Takiguchi et al., 1980; Stapley and Woodruff, 1982;

Takiguchi et al., 1983). MLs are categorized as avermectins or milbemycins. Together, they have a common 16-member macrocyclic lactone ring that is fused with spiroketal and benzofuran functions in a three-dimensional arrangement (Campbell, 1989; Prichard & Geary, 2019).

Commercially available avermectins include abamectin, ivermectin, emamectin, eprinomectin, doramectin and selamcectin, while milbemycins include nemadectin, moxidectin and milbemycin oxime (Figure 1.15). Ivermectin, emamectin and eprinomectin are chemical derivatives of abamectin. On the other hand, moxidectin was derived chemically from nemadectin. Collectively, these compounds were developed and approved for use against nematodes and arthropods in the 1980s.

Source: Prichard and Geary, 2019, p. 70.

Fig 1.15 Chemical structures of commercially available macrocyclic lactones

The mode of action of the MLs involve selective binding of the drug with high affinity to glutamate-gated chloride channels (GluCls) that are expressed in the pharyngeal muscles and neurons of nematodes and arthropods. Consequently, this increases the influx of Cl- ions, resulting in hyperpolarization and paralysis of the pharyngeal and body wall musculature (Cully et al., 1994; Wolstenholme & Rogers, 2005). It is noteworthy that GluCls are absent in humans, making it a valuable target for anthelmintic therapy. Avermectins anthelmintic activity are not limited to GluCls, but involve interaction with nAChRs, 4-aminobutyric acid (GABA) and glycine receptors where diverse pharmacological effects were exhibited. Krause et al. (1998) reported that ivermectin acted as a positive allosteric modulator on the vertebrate α7 nicotinic acetylcholine receptor. In another study ivermectin was shown to act as an irreversible agonist 54 and positive allosteric modulator at higher (> 0.3 µM) and lower (30 nM) concentrations respectively on recombinant human glycine receptors (Shan et al., 2001). Avermectins have also had antagonistic effects on nAChRs, and GABA receptors expressed in parasitic nematodes somatic muscle (Holden-Dye & Walker, 1990; Puttachary et al., 2013 Abongwa et al., 2016).

Benzimidazole (BZs)

The benzimidazoles are a group of anthelmintic drugs that are widely used in the treatment of parasitic nematode infections worldwide. The first of this class, thiabendazole, was discovered in 1961 which was subsequently followed by albendazole, mebendazole and flubendazole (Figure 1.16.). The mode of action of this drug class was initially considered to be involved in the inhibition of parasitic nematodes metabolic enzymes. Prichard (1970) reported that the mode of action of thiabendazole was to inhibit malate dehydrogenase in H. contortus. In a later study, Tejada et al. (1987) revealed that four benzimidazole anthelmintics, namely, mebendazole, parbendazole, albendazole and thiabendazole had inhibitory effects on purified extracts of mitochondrial and cytoplasmic malate dehydrogenase in Moniezia expansa, Fasciola hepatica and Ascaris suum. However, later experiments established that the mode of action of benzimidazoles, is to selectively bind with high affinity to nematode β-tubulin, thus inhibiting microtubule formation. Consequently, this results in the disruption of intestinal cell structure, starvation, inhibition of egg production and subsequent death of the parasite (Lacey, 1990). The effects of these drugs however are generally slower when compared to nicotinic agonists.

Moreover, they have been proven to be efficacious and possess broad spectrum activity.

Source: McKellar & Scott, 1990, p. 227

Figure 1.16 Chemical structures of benzimidazole anthelmintics, namely, (A) thiabendazole, (B) mebendazole, (C) Albendazole and (D) Flubendozole.

Anthelmintic resistance

Anthelmintic resistance poses a serious threat to animal and human health and global food security. It is defined as a phenomenon where an anthelmintic losses efficacy against a population of a species of parasites at concentrations that once rendered them susceptible

(Prichard et al., 1980; Coles et al., 2006). Anthelmintic resistance can be conferred through a selection process, where a population of parasites survive drug treatment at a concentration that kills others within the population. Parasites that survive possess a resistant phenotype, while those that die, possess a susceptible phenotype. Consequently, genes responsible for resistance in the surviving parasites are passed on to the next generation where they are transmitted to another host. There are cases where a single gene may confer resistance, while in others multiple genes might be involved. Initially, these resistant genes are rare in the population, or may have appeared by random mutations (Geary et al., 2012). However, as selection continues, the 56 proportion or frequency of the resistant genes increases, thus increasing the proportion of resistant parasites (Geary et al., 2012).

For decades, anthelmintic drugs have been widely used as the primary mode for successfully controlling parasitic nematode infections. However, use of these drugs has led to increase in the incidence of anthelmintic resistance worldwide, thus stimulating numerous research that are aimed at developing new control measures. Resistance has been reported in several phyla of helminths that affect cattle, sheep, goats, horses, pigs and humans. The incidence of resistance in human parasites, however, are more difficult to estimate since field data is harder to obtain. Drug classes associated with resistance include the macrocyclic lactones, benzimidazoles, imidazothiazoles/tetrahydropyrimidines, aminoacetonitrile derivatives and spiroindole (Sangster & Gill, 1999; Kaplan, 2004; Scott et al., 2013; Sangster et al., 2018).

Resistance to anthelmintics can be classified as side resistance or cross resistance. Side resistance involves the development of resistance to one anthelmintic of a certain class, which leads to resistance among all the drugs of that same class due to shared similarities in mode of action

(Sangster, 1999). Cross resistance on the other hand occurs when a parasite develop resistance to more than one anthelmintic class, providing that the two drug classes have similar therapeutic targets (Sangster, 1999; Torres-Acosta et al., 2012).

The earliest official report of anthelmintic resistance was in 1957, where H. contortus resistance to phenothiazine in treated sheep was discovered (Drudge et al., 1957). Within the next 10 years following the official report, resistance was found regularly in sheep parasites followed by cattle and horse nematodes (Jackson et al., 2006). Resistance has been reported in countries such as New Zealand, Australia, Italy, South Africa and Switzerland where there are large sheep production systems (Overend et al., 1994; Leathwick et al., 2001; Schnyder et al., 57

2005; Cringoli et al., 2007). Drugs involved in high levels of resistance are fenbendazole, albendazole and ivermectin, while moderate to low resistance have been seen with levamisole.

Moxidectin resistance has also been observed at a high rate on numerous livestock farms

(Prichard & Geary, 2019). Resistance is also seen in monepantel and derquantel although it has not been widespread (Sangster et al., 2018). High levels of anthelmintic resistance have also been reported in 13 countries in Africa, with South Africa been the leader with the highest levels of H. contortus resistance to all classes of anthelmintics (Anon, 2007). Despite the numerous cases of anthelmintic resistance globally, the mechanisms of resistance have not been fully elucidated. Currently, there are several proposed mechanisms of resistance. This includes: (i) changes in the molecular target, or receptor binding site due to mutations; (ii) reduction in the number of receptors (iii) a change in metabolism that inactivates or remove drug; (iv) a change in drug distribution in the parasite, thus preventing the drug from accessing its site of action

(Sangster & Gill, 1999; Wolstenholme et al., 2004; Gilleard, 2006; Jabbar et al., 2006; Martin &

Robertson, 2007; James et al., 2009).

Resistance can be delayed through several management strategies. This includes: (i) combination therapy with drugs from different classes; (ii) selective treatment of individuals that require treatment on the basis of their parasitism, rather than treating the whole population (iii) eliminating suboptimal dosing and over-use of the same anthelmintic class (iv) keeping some parasites in untreated refugia; and (v) identifying new drug targets and synthesizing novel anthelmintics that target the newly discovered site. Collectively, anthelmintic resistance might be resolved once these strategies are adopted universally and put into practice. This has been proven based on evidence from New Zealand where reversion in macrocyclic lactone and levamisole resistance have been observed (Leathwick et al., 2015). 58

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Williamson, S.M., Robertson, A.P., Brown, L., Williams, T., Woods, D.J., Martin, R.J., Sattelle D.B., Wolstenholme, A.J. 2009. The nicotinic acetylcholine receptors of the parasitic nematode Ascaris suum: formation of two distinct drug targets by varying the relative expression levels of two subunits. PLoS Pathog, 5, e1000517. doi: 10.1371/journal.ppat.1000517.

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CHAPTER 2. CHOLINERGIC RECEPTORS ON INTESTINE CELLS OF ASCARIS SUUM AND ACTIVATION OF NACHRS BY LEVAMISOLE

Mark McHugha, Paul Williamsb, Saurabh Vermab, Jo Anne Powell-Coffmana, Alan P.

Robertsonb, Richard J. Martinb*

a Department of Genetics, Development and Cell Biology, Iowa State University, Ames,

Iowa USA

b Department of Biomedical Sciences, College of Veterinary Medicine, Iowa State

University, Ames, Iowa, USA

*Corresponding author: [email protected]

Modified from a manuscript published in the International Journal for Parasitology:

Drugs and Drug Resistance

2.1 Abstract

Cholinergic agonists, like levamisole, are a major class of anthelmintic drug that are known to act selectively on nicotinic acetylcholine receptors (nAChRs) on the somatic muscle and nerves of nematode parasites to produce their contraction and spastic paralysis. Previous studies have suggested that in addition to the nAChRs found on muscle and nerves, there are nAChRs on non-excitable tissues of nematode parasites. We looked for evidence of nAChRs expression in the cells of the intestine of the large pig nematode, Ascaris suum using RT-PCR and RNAscope in situ hybridization and detected mRNA of nAChR subunits in the cells. These subunits include components of the putative levamisole receptor in A. suum muscle: Asu-unc-38,

Asu-unc-29, Asu-unc-63 and Asu-acr-8. Relative expression of these mRNAs in A. suum 80 intestine was quantified by qPCR. We also looked for and found expression of G protein-linked acetylcholine receptors (Asu-gar-1). We used Fluo-3 AM to detect intracellular calcium changes in response to receptor activation by acetylcholine (as a non-selective agonist) and levamisole (as an L-type nAChR agonist) to look for evidence of functioning nAChRs in the intestine. We found that both acetylcholine and levamisole elicited increases in intracellular calcium but their signal profiles in isolated intestinal tissues were different, suggesting activation of different receptor sets. The levamisole responses were blocked by mecamylamine, a nicotinic receptor antagonist in A. suum, indicating the activation of intestinal nAChRs rather than G protein-linked acetylcholine receptors (GARs) by levamisole. The detection of nAChRs in cells of the intestine, in addition to those on muscles and nerves, reveals another site of action of the cholinergic anthelmintics and a site that may contribute to the synergistic interactions of cholinergic anthelmintics with other anthelmintics that affect the intestine (Cry5B).

Keywords

Levamisole; nicotinic acetylcholine receptors; intestine; calcium signaling

2.2 Introduction

Parasitic infections by soil-transmitted helminths (STHs: Ascaris, Trichuris and hookworm) are a major medical and public health concern in many developing countries. The frequency of these infections is staggering, with estimates of approximately 1.5 billion humans being infected globally (WHO, 2017); over the world, the high morbidity reduces worker productivity by 6.3 million Disability Adjusted Life Years (DALYs) per year. Poverty is correlated with the levels of these infections that degrades worker health, worker output and school performance in children. There is also impairment of the immune system, leading to exacerbation of HIV/AIDS and increased susceptibility to other illnesses such as tuberculosis and malaria infection (Fincham et al., 2003; Le Hesran et al., 2004; Brooker et al., 2007; 81

Alexander and De, 2009). In livestock, STHs cause lost food production and reduced economic returns that contribute further to poverty (De Silva et al., 2003; Puttachary et al., 2013).

In the absence of adequate sanitation and effective vaccines, control of these infections relies on three major classes of anthelmintic drugs: benzimidazoles like albendazole, the macrocyclic lactones like ivermectin, and the nicotinic cholinergics like levamisole and pyrantel.

Pyrantel and levamisole act as agonists that selectively gate nicotinic acetylcholine receptor

(nAChR) channels on muscle cells of the parasite (Martin and Robertson, 2010; Abongwa et al.,

2017). Opening of muscle nAChRs produces depolarization, spastic paralysis and consequent expulsion of the parasite from the host (Aubry, 1970; Aceves, 1970; Martin and Robertson,

2007). Frequent use of these anthelmintics has led to the emergence of widespread drug resistance in animal parasites (Prichard, 1994) and there is concern about the development of resistance in human parasites. The limited number of anthelmintic drugs and the development of resistance drives the need to determine how existing anthelmintic act, to understand and control resistance and develop novel drugs and combinations.

nAChRs, the target sites of cholinergic anthelmintics, are pentameric membrane proteins comprised of five subunits that surround a central cation-permeable pore (Devillers-Thiery et al.,

1993; Unwin, 1993). Combinations of different nAChR subunits leads to the formation of heteromeric receptors on muscle cells that give rise to a diversity of receptor subtypes, each with different pharmacology and anthelmintic sensitivities (Williamson et al., 2009; Buxton et al.,

2014; Verma et al., 2017). In vertebrates and the parasitic nematode Brugia malayi, nAChR subunit expression has been shown to be in both excitable cells, like nerve and muscle, and in non-excitable cells. Examples of nAChRs on non-excitable cells in mammals include: human skin keratinocytes (Grando, 1995), human bronchial epithelial and endothelial cells (Wang et al., 82

2001), human vascular endothelial cells (Macklin et al., 1998), mouse intestine and intestinal crypt-villus organoids that lack nerve (Takashi et al., 2018). In Brugia malayi, the presence of nAChR message has been reported in the developing embryo and spermatozoa (Li et al., 2015) but physiological evidence of activation of nAChRs in non-excitable cells in nematodes is lacking.

Of particular interest to us here, is the presence of functioning nAChRs on cells other than muscles and nerves. These paracrine nAChRs on non-excitable cells in nematode parasites may be activated by cholinergic anthelmintics and play a role in their effects on the parasites.

This interest was provoked by the potentiating effect of cholinergic anthelmintics on the action of Cry5B, a pore-forming peptide Bt toxin that acts on the intestinal tract of nematodes (Hu et al., 2010; 2018). These observations suggest the presence of nAChRs within the intestine that are also the site of action of the cholinergic anthelmintic drugs and that a combination of Cry5B and cholinergic anthelmintics could be advantageous.

In this paper we explore the presence of nAChRs in the nematode intestine. The nematode intestine is important for enzymatic digestion and nutrient absorption (McGhee, 2007;

Yin et al., 2008). In addition, other processes such as ion transport (Nehrke, 2003), defense against environmental toxins (Park et al., 2001; Rosa et al., 2015) and innate immunity to microbial infection (Rosa et al., 2015; Schulenburg et al., 2004) are present in the intestine of nematodes.

We used Ascaris suum for our studies (Martin, 1993). This parasite is closely related to

Ascaris lumbricoides seen in humans (Boes and Helwigh, 2000; Nejsum et al., 2005). We used

RT-PCR, quantitative Real-time PCR (qPCR), and RNAscope to identify and quantify, in the muscle and intestine of adult female A. suum, the relative expression of message of the nAChR 83 subunits that constitute the putative levamisole receptor, namely: Asu-unc-38, Asu-unc-29, Asu- unc-63 and Asu-acr-8. RNAscope is an in-situ hybridization technique that uses a novel signal amplification method that allows for the detection, visualization and localization of target RNAs as punctate dots in individual cells (Wang et al., 2012; Anderson et al., 2016). Our observations indicate the presence of nAChRs in the intestine. Furthermore, we used calcium imaging to show that application of acetylcholine and levamisole to the intestine transiently increases cytoplasmic calcium. Our findings revealed evidence of functioning nAChRs in the intestine that may be exploited further by existing cholinergic anthelmintics or novel therapeutic combinations.

2.3 Materials and Methods

2.3.1 Collection and maintenance of A. suum worms

Adult female A. suum worms were collected from the JBS Swift and Co. pork processing plant at Marshalltown, Iowa. Worms were maintained in Ascaris Ringers Solution (ARS: 13 mM

NaCl, 9 mM CaCl2, 7 mM MgCl2, 12 mM C4H11NO3/ Tris, 99 mM NaC2H3O2, 19 mM KCl and 5 mM glucose pH 7.8) at 32 °C for 24 hrs. to allow for acclimatization prior to use in experiments. The worms were used the next day for experiments.

2.3.2 Histological preparation of A. suum

For histological analysis of the morphology of A. suum, 1 cm of an adult female worm was cut transversely, ~3 cm caudal to the pharyngeal region. Subsequently, the sample was washed in phosphate buffered saline (PBS) and immediately fixed in 10% neutral-buffered formalin for 24 hrs. at room temperature. The fixed sample was dehydrated and infiltrated with paraffin, followed by manual embedding into paraffin wax blocks. Tissue sections of 5 µm thickness were made using a microtome after hardening of the paraffin wax mold, and subsequently mounted on Superfrost® Plus microscope slides (Fisher Scientific Pittsburgh, PA,

USA). The section was subjected to hematoxylin and eosin staining and observed with an 84

Olympus BX53 Microscope® (Olympus America, Inc., Center Valley, PA, USA), equipped with an Olympus DP73 (Olympus America, Inc., Center Valley, PA, USA) camera which was used for capturing images using the cellSensTM Imaging Software version 1.12 (Olympus

Corporation of the Americas, Waltham, MA, USA).

2.3.3 Intestinal tissue and muscle bag preparation

Dissection was conducted on adult A. suum females (n=5) by cutting a 2 cm section from the anterior region of the worm, ~3 cm caudal to the pharyngeal region. The resulting section was cut along one of the lateral lines and pinned cuticle side down onto a 35 x 10 mm Petri-dish lined with Sylgard, to form a muscle flap, thus exposing the intestine and muscle bags. The intestine was gently removed using fine forceps. Next, the body wall flap with exposed muscle bags was washed with autoclaved Ascaris perienteric fluid (APF: 23 mM NaCl, 110 mM Na acetate, 24 mM KCl, 6 mM CaCl2, 5 mM MgCl2, 11mM glucose, 5 mM HEPES, pH 7.6) to remove fragments of the intestine. The APF solution was then replaced with collagenase solution for a period of 5 min. at room temperature to facilitate the separation of muscle bags from the body wall of the worm. After collagenase treatment, the muscle flap was washed with APF, and muscle bags were separated from the body wall with fine forceps, and collected with a micropipette. Both intestine and muscle bag tissue were immediately snap-frozen in liquid nitrogen and stored at – 80°C until further use.

2.3.4 A. suum cDNA synthesis and RT-PCR detection of nAChR subunits

Muscle bags and intestinal tissue of A. suum were homogenized separately in 1 ml of

Trizol reagent using a mortar and pestle, followed by total RNA extraction according to the

Trizol Reagent protocol (Invitrogen/Life Technologies, Carlsbad, CA, USA). One microgram (1

µg) of total RNA from each tissue was treated with DNase I (Quanta Biosciences, Inc.,

Gaithersburg, MD, USA) for the removal of residual genomic DNA. This was followed by 85 reverse transcription (RT) using qScriptTM Flex cDNA Synthesis kit (Quanta Biosciences, Inc.,

Gaithersburg, MD, USA) following the manufacturer’s protocol. PCR was conducted to detect the presence of Asu-unc-38 (EU053155.1), Asu-unc-63 (KY654348.1), Asu-unc-29

(EU006073.1) and Asu-acr-8 (KY654347) using primers that were designed for targeting transmembrane 1 to transmembrane 4 (TM1-TM4) regions of each gene. Asu-gapdh

(AB058666.1) was used as a reference gene. We also looked for expression of the G protein- linked acetylcholine receptor, Asu-gar-1 (FJ609743.1) in the intestine of adult female worms.

Asu-gar-1 is alternatively spliced with two isoforms: Asu-gar-1a (1,956 bp) and Asu-gar-1b

(1,875 bp). We aligned the nucleotide sequences of both isoforms a and b using the Clustal

Omega multiple sequence alignment tool and designed receptor-specific primers targeting the end of transmembrane 2 and downstream transmembrane 5 where both isoforms had 100% similarity. Again, Asu-gapdh was used as a reference gene. Primers for the other genes namely,

Asu-unc-29, Asu-unc-63, Asu-unc-38, Asu-acr-8 and Asu-gapdh are presented in Table S2.1. The cycling conditions for PCR were an initial denaturation for 2 min at 95 °C, followed by 35 cycles of 95 °C for 30 sec, 54 °C for 35 sec, 72 °C for 45 sec, and a final extension at 72 °C for 5 min.

PCR products were then separated on a 1% Agarose ethidium bromide gel, followed by visualization and sequencing to confirm the identity of the subunits.

2.3.5 Analysis of mRNA levels by Quantitative Real-time PCR

To quantify the relative mRNA transcript levels of each subunit in the intestine and muscle bags, cDNA was synthesized from 1 µg of three adult female A. suum total RNA samples

(n = 3) that were extracted for use in PCR experiments. Quantification standards were generated by pooling equal volumes of cDNA from each worm sample followed by serial dilutions. Target genes with fragments ranging from 150 to 200 bp between TM3 and TM4 regions were amplified by qPCR from each cDNA sample. This was also done for the reference gene Asu- 86 gapdh. All primers for qPCR are presented in Table S2.2. The real-time PCR reaction mixture consisted of 1 µl of cDNA template, 1 µl of primer mix (500 nM [each]), and 10 µl of

SsoAdvancedTM Universal SYBR® Green Supermix (Bio-Rad), with the final volume made up to 20 µl with Nuclease-free water. The cycling conditions included an initial denaturation for 30 sec at 95oC, 40 cycles of 95oC for 15 sec, 54oC for 20 sec, 65oC for 5 sec and a final melting curve step. Cycling was performed using a CFX 96 real-time system (Bio-Rad, Hercules, CA) and transcript quantities were derived by the system software, using the generated standard curves. mRNA expression levels for each subunit (Asu-unc-38, Asu-unc-29, Asu-unc-63 and Asu- acr-8) was estimated relative to the reference gene (Asu-gapdh) using the Pfaffl Method. The qPCR experiments were repeated 3 times for each gene (all subunit mRNA quantifications were performed in triplicate for each worm’s muscle bag sample and intestinal tissue sample: 3 biological replicates each with 3 technical replicates). Statistical analysis was done using the one-way analysis of variance (ANOVA), P < 0.05 with GraphPad Prism 5.0 (Graphpad

Software, Inc., La Jolla, CA, USA). The data are presented as relative expression levels in mean

± SEM X1000 units for each nAChR subunit relative to Asu-gapdh in muscle bags and intestinal tissue. Comparisons were made using Tukey’s post-hoc test.

2.3.6 Preparation of Formalin-Fixed Paraffin-Embedded (FFPE) tissue for RNAscope ISH

Cross sectional dissections of 1 cm was made from adult female A. suum worms (n = 3)

~4 cm below the pharyngeal region. Each dissected worm sample was immediately placed in fresh 10% neutral-buffered formalin (NBF) for 24 h at room temperature, followed by dehydration with paraffin, and manual embedding into paraffin wax blocks. Tissue sections of 5

µm thickness were then made using a microtome, and subsequently mounted on Superfrost®

Plus slides (Fisher Scientific Pittsburgh, PA, USA). 87

2.3.7 RNAscope in situ hybridization Assay

RNAscope in situ hybridization (RNAscope ISH) was conducted on FFPE tissue sections using the RNAscope® 2.0 HD Assay (Advanced Cell Diagnostics Inc., Hayward, CA, USA) according to the manufacturer’s instructions. FFPE tissue section slides were deparaffinized by incubating for 60 min at 60°C, followed by submersion in xylene for 5 min, incubation in 100% ethanol for 1 min and then air dried. Subsequently, each section was pre-treated for cell permeabilization and target RNA access, using the RNAscope pretreatment reagents. The individual target probes Asu-unc-38 (Catalog # 480948), Asu-unc-29 (Catalog #480938) and

Asu-unc-63 (Catalog # 480951) along with negative (dapB, targets bacterial gene) and positive

(Asu-gapdh) control probes were then hybridized for 2 h at 40°C, followed by RNAscope amplification, using several hybridization buffers (Amp 1-6). Subsequently, each slide was washed with wash buffer for 2 min at room temperature. Fast red detection was performed, followed by DAB chromogenic detection and counterstaining with hematoxylin. Stained slides were examined with an Olympus BX53 Microscope® (Olympus America, Inc., Center Valley,

PA) with pink punctate dots representing positive expression of mRNA transcript for each subunit. All probes were designed and synthesized by Advanced Cell Diagnostics.

2.3.8 Subcellular quantification of mRNA punctate dot analysis

After the RNAscope ISH assay, mRNA transcript (pink punctate microdots) abundance for each of the investigated subunits was quantified across all intestinal tissue visible in the cross sections and cell regions divided into basolateral, nuclear, peri-nuclear, central and apical areas for each of the intestines of three worm (n = 3), Fig 2.4A. The data were expressed as mean density of mRNA transcripts/µm2. We also quantified the density of the subunit mRNAs dots in the basolateral (B) or central (C) regions of the columnar intestinal cells, Fig 2.4A. 88

For accurate quantification of punctate dots in the intestine of each worm, the intestine was divided into several photographed frames with each image captured at 200 x magnification.

These frames covered an area of the intestine of ~10,000 µm2. Each cross-sectional slide of the worms was viewed with an Olympus BX53 Microscope® (Olympus America, Inc., Center

Valley, PA, USA), equipped with an Olympus DP73 camera (Olympus America, Inc., Center

Valley, PA, USA) that was connected to the microscope. The camera and microscope were connected to a computer where images were transferred and displayed using the cellSensTM

Imaging Software version 1.12 (Olympus Corporation of the Americas, Waltham, MA, USA).

Caution was taken not to capture overlapping regions. The total area of the intestinal sections in each frame was acquired using the HaloTM Image Analysis Software version 2.0.1145.19

(Indica Labs, Corrales, NM, USA). Each image frame was then analyzed manually to count the number of punctate dots for the subunits using the Windows 10 Paint Desktop App.

The total number of dots were tabulated for each frame and cell region in Microsoft

Excel. To calculate the density of each subunit distribution in the intestine cells, the total number of punctate dots in each frame was divided by the measured area of the intestine (µm2). The data was analyzed using GraphPad Prism 5.0 (Graphpad Software Inc., La Jolla, CA, USA). We used one-way ANOVA followed by the Tukey multiple comparison test, to determine statistical differences (P < 0.05) between the densities of the subunits for each intestinal cell region.

Results were displayed as mean ± SEM.

2.3.9 Intestinal Calcium Imaging

A 2 cm section of intestine was collected as described earlier with fine forceps and cut open to make a flap. Subsequently, the intestinal flap was placed onto a coverslip (24 x 50 mm) and pinned using a slice anchor (26 x 1mm x 1.5mm grid, Warner Instruments, Hamden, CT), immersed in Ascaris Perienteric Fluid with no added CaCl2 in a laminar flow chamber (Warner 89

RC26G, Warner Instruments, Hamden, CT). The calcium concentration of APF with no added

CaCl2 was measured and found to be < 100 µM. Fluo-3AM loading was achieved by incubating the intestine in APF solution with no added CaCl2 containing 5 µM Fluo-3AM and 10% Pluronic

F-127 (10% v/v) for 1 h with the chamber connected to a Dual Automatic Temperature

Controller (Warner Instruments, Hamden, CT). The chamber temperature was maintained between 34 and 36°C to allow cleavage of the ester group by endogenous non-specific esterases.

After 1 h, the excess Fluo-3AM solution was discarded, and the sample was incubated in APF

2+ containing 500 µM CaCl2 for an additional 15 min at 34-36°C to promote Ca loading into the intestinal cells. All incubations were done in the absence of light to prevent degradation of the fluorescent dye. After incubation the sample was continuously perfused with APF containing

500 µM CaCl2. All solutions were delivered to the chamber under gravity feed through solenoid valves controlled using a VC-6 six channel Valve Controller (Warner Instruments, Hamden, CT) through an inline heater set at 37°C (Warner Instruments, Hamden, CT) at a rate of 1.5mL/min.

At the start of all experiments, samples were exposed to APF containing 500µM CaCl2 wash under blue light for a minimum of 3 min to promote settling and equilibration of the fluorescent signal and to monitor for any spontaneous Ca2+ signaling. Intestinal tissues were then bathed in 30 µM acetylcholine, or 30 µM levamisole, or 30 µM acetylcholine followed by 30

µM levamisole after a 5 min wash after a drug application. For the antagonist experiment, samples were exposed to either 10 µM mecamylamine (antagonist) alone, a combination of 10

µM mecamylamine and 30 µM levamisole or 30 µM levamisole followed by a combination of

10 µM mecamylamine and 30 µM levamisole if an increase in Ca2+ was observed. 30 µM acetylcholine, 30 µM levamisole and 10 µM mecamylamine applications were made within APF containing 500 µM CaCl2. All samples were then tested with 10 mM CaCl2 after drug 90

applications as a positive control and for response amplitude comparisons. For 10 mM CaCl2 a total of 689 regional measurements were obtained from 12 intestinal preparations (n = 6 adult female worms). For acetylcholine, 280 regional measurements were analyzed from 7 intestinal preparations (n = 4 adult female worms) and for levamisole, a total 241 regional measurements were analyzed from 5 intestinal preparations (n = 4 adult female worms). Mecamylamine had

300 regional measurements from 5 intestinal preparations (n = 3 adult female worms) and levamisole + mecamylamine had 230 recordings from 4 intestinal preparations (n = 4 adult females). A region consisted of a square area of 50 µm x 50 µm of adjacent cells.

All recordings were performed on a Nikon Eclipse TE3000 microscope (20X/0.45 Nikon

PlanFluor objective), fitted with a Photometrics Retiga R1 Camera (Surrey, BC, Canada). Light control was achieved using a Lambda 10-2 two filter wheel system with shutter controller

(Lambda Instruments, Switzerland). Filter wheel one was set on a green filter between the microscope and camera. Filter wheel two was set on the blue filter between a Lambda LS Xenon bulb light box which delivered light via a fiber optic cable to the microscope (Lambada

Instruments, Switzerland). Blue light emission was controlled by using a shutter. Minimal illumination exposure was used to prevent photo bleaching.

All Ca2+ signal recordings were acquired and analyzed using MetaFluor 7.10.2 (MDS

Analytical Technologies, Sunnyvale, CA) with exposure settings at 250 ms with 2x binning.

Maximal Ca2+ signal amplitudes (ΔF) were calculated using the equation F1-F0/F0 x 100, where

F1 is the fluorescent value and F0 is the baseline value. All F0 values were determined as being the value at the time any stimulus was applied to the sample for all traces analyzed. All pictures were taken using Occular 2.0.1.496 (Digital optics, Auckland, New Zealand). Exposure settings 91 were 150 ms with 2x binning. Statistical analysis was performed using two-tailed unpaired student t-tests in GraphPad Prism 5.0 (Graphpad Software, Inc., La Jolla, CA, USA).

2.4 Results

2.4.1 Transverse section of Ascaris suum

Fig. 2.1 shows a hematoxylin and eosin stained transverse section through an Ascaris suum female worm, 1 cm anterior to the vulva. The section shows the cuticle (C) that surrounds the worm, beneath which lies the hypodermis (H). The muscle tissue includes the contractile spindles (S) and arms (A) that connect the spindles to the nerve cords and the bag region (B).

The dorsal and ventral halves of the animal are separated by two lateral lines (L), and the intestine (I). Canals (Cn) that form the perienteric space are found between the intestine and bag regions of the muscle cells; they provide a mechanism for fluid transport along the parasite.

In addition, nutrients may be absorbed from the intestine and then moved, through the perienteric canals (Cn). The muscle cells are innervated by excitatory or inhibitor motor neurons of the dorsal (DN) or ventral (VN) nerve cords that are at the end of the arms.

2.4.2 nAChR subunit mRNA expression in A. suum muscle and intestinal cells

Rosa et al. (2014) used RNA-seq to indicate the presence of nAChR subunit message in the intestine of A. suum. We isolated mRNA from explanted intestine and muscle tissue from 5 individual adult Ascaris and used RT-PCR to identify the presence of specific nAChR subunits.

We looked for and detected expression of Asu-unc-29, Asu-unc-63, Asu-unc-38 and Asu-acr-8, in both intestinal cells and muscle bags. These data are shown in Fig. 2.2. The brightest bands in intestine cells were Asu-unc-38 and Asu-acr-8; Asu-unc-29 bands were in contrast fainter than the others.

2.4.3 Differential expression of nAChR subunits in muscle bags and intestinal tissue

Expression of the putative L-type nAChR subunits in the muscle bag regions and the intestinal tissue prompted us to measure the relative mRNA expression levels of each subunit to examine the difference. Using quantitative real time PCR (qPCR), we compared mRNA expression levels of each subunit in the intestine and muscle bag of the parasite relative to a reference gene, Asu-gapdh.

Our analysis of the muscle bag region showed that mean mRNA expression levels for

Asu-unc-38 was 6-fold higher than that of Asu-unc-63, and approximately 30 times higher than

Asu-acr-8 and Asu-unc-29 (Fig. 2.3 A). Asu-unc-63 mRNA levels were 5-fold greater than Asu- unc-29 and 3.5-fold higher than Asu-acr-8, Fig. 2.3A. The much higher expression levels of Asu- unc-38 is interesting and suggests that it may contribute to more than one type of nAChR.

When we looked at expression levels of mRNA in the intestine, we found them to be much lower than in the muscle, Fig. 2.3B. The muscle Asu-unc-38, for example, was nearly 7x higher than the intestine Asu-unc-38. Although the mean mRNA expression levels of intestine

Asu-unc-38 appeared greater than Asu-acr-8, which in turn appeared greater than Asu-unc-29 and

Asu-unc-63, the sample-to-sample variation was high, lessening confidence that the values are significantly and consistently different. Nonetheless, the presence of mRNA for the subunits of the putative L-type nAChR in the intestine suggests the presence of levamisole receptors there.

2.4.4 RNAscope reveals heterogeneous subcellular distribution of intestinal nAChR subunit mRNAs

We used RNAscope in situ hybridization (ISH) to localize the distribution of nAChR subunit mRNAs within the intestinal cells. Each individual 1-2µm pink dot, represents an individual RNA molecule and its location within cells. Fig. 2.4 illustrates representative examples of our RNAscope data. 93

The intestine is composed of columnar cells that extend from the absorptive brush border on the apical edge, with nuclei located near the basolateral border, Fig. 2.4A. For RNAscope experiments, we examined transverse histological sections of the intestine for each of the four probes in three individual adult female worms. Fig. 2.4B shows the results from a Bacillus dapB probe, which serves as a negative control; note the absence of any pink punctate dots which would mark mRNA single message strands. Fig. S2.1 also shows results of Asu-gapdh positive control; attention should be drawn to the presence of numerous punctate dots throughout the tissue. The presence of Asu-unc-29 (Fig. 2.4C), Asu-unc-63 (Fig. 2.4D), Asu-unc-38 (Fig. 2.4E) and Asu-acr-8 (Fig. 2.4F) mRNAs in the columnar cells of the intestine was also seen in all sections examined. Intriguingly, the distribution patterns of the mRNA transcripts for the subunits were heterogeneous. Asu-unc-29 and Asu-unc-63 was characterized by sparse distributions of microdots that were more focused in the nuclear and basolateral regions of the intestine, (see arrows: Fig. 2.4C & 2.4D). In contrast, Asu-unc-38 and Asu-acr-8 mRNAs were distributed along the length of the columnar cells (Fig. 2.4E & 2.4F) with a large number of microdots within the central region. These observations from the RNAscope studies, showing the presence of the different nAChR subunit message, coupled with our RT-PCR results, provided further molecular evidence of levamisole-sensitive, L-type nAChRs, in Ascaris suum intestinal cells.

2.4.5 Subcellular distribution of intestinal nAChR subunit mRNAs

The differences in the subcellular localization pattern of each subunit mRNAs in our

RNAscope assay led us to quantify those differences. We analyzed the mean density of each subunit mRNA transcripts/µm2 within the intestinal cells and in the basolateral (B) and central

(C) sub-cellular regions of individual cells where the heterogeneous distribution patterns were observed, Figs 2.5A. We found that the mean density of Asu-unc-38 mRNA was significantly 94 greater in the columnar cells of the intestine, compared to the mean density of Asu-unc-29 and

Asu-unc-63; and the mean density of Asu-acr-8 was somewhere between that of Asu-unc-38 and

Asu-unc-29 (Fig. 2.5A). Results from our quantification of the whole intestine revealed that the

RNAscope data was consistent with our earlier qPCR findings. Asu-unc-38 mRNAs were more abundant in the intestine, compared to Asu-unc-29 and Asu-unc-63.

RNAscope also revealed a lower density of all subunit mRNAs in the basolateral region of the columnar cells of the intestine compared to the mean distribution throughout the cell, Fig.

2.5B; but the Asu-unc-38 transcripts were still the most abundant in the basolateral region when compared to Asu-acr-8 (P < 0.05), Asu-unc-29 (P < 0.01) and Asu-unc-63 (P < 0.001).

The mRNA transcripts for Asu-unc-38 and Asu-acr-8 in the central region of the intestine columnar cells were more abundant than the transcripts of Asu-unc-29, and Asu-unc-63, Fig.

2.5C. Collectively, these findings of subunit mRNA distribution heterogeneity in the basolateral and central regions, suggest different concentrations of nAChR subunits in different regions of the columnar cells and consequently the presence of different nAChR subtypes in different regions of the intestine columnar cells.

2.4.6 Acetylcholine and levamisole stimulate calcium signals in Ascaris intestine

Having determined that nAChR subunit mRNAs were expressed in the intestine of

Ascaris suum, we sought evidence of functional responses to acetylcholine and levamisole. To do this we looked for intracellular calcium signals as important markers of activation in the intestinal cells.

To check for the presence of autofluorescence in the intestine, dissected lengths of each worm intestine incubated in Fluo-3AM-free solutions were compared with intestine incubated in

5 µM Fluo-3AM containing 10% Pluronic F-127 (v/v). The micrographs of bright-field (Fig.

3.5A, left image) and 5 µM Fluo-3AM treated intestinal tissues (Fig. 2.6A, right image) showed 95 the presence of individual cells as fluorescent spots arranged in groups or tower like microvilli.

No equivalent fluorescence of individual cells was observed in the untreated tissue (Fig. 2.6A, center image). To ensure that changes in Ca2+ could be measured in Ascaris intestine cells, we applied 10 mM CaCl2. Fig 2.6B shows a representative result. Following application of CaCl2, a detectable Ca2+ signal was observed that, on average, peaked after 160 ± 1.7 sec, with an average increase of 28 ± 0.88% in Fluo-3 fluorescence, which declined shortly after CaCl2 was removed

(Fig. 2.6B & C). We observed no distinct changes in the Ca2+ signal in samples continuously perfused with 500 µM CaCl2 APF (Fig. 2.6C, APF vs CaCl2).

To measure acetylcholine generated changes in Ca2+ signals, we applied 30 µM acetylcholine to the intestine. Acetylcholine (30 μM) generated unique Fluo-3 responses which were typically characterized as a slow and steady rise in Ca2+ amplitude, which continued to increase even after removal of acetylcholine (Fig. 2.6D.) Unlike CaCl2, the acetylcholine responses took significantly longer to reach a peak after application (554 ± 17.49 s) and had a slower decline after peak had been reached (Fig. 2.6D & F). The overall increases in Fluo-3 fluorescence for acetylcholine were smaller than for the 10 mM CaCl2 test and averaged an increase of 12 ± 0.41% (Fig. 2.6G). The acetylcholine responses also showed greater fatigue with subsequent applications of acetylcholine being smaller than the first responses.

Acetylcholine is a non-selective cholinergic agonist and can activate a range of acetylcholine receptors including G-protein coupled receptors and different types of nAChRs. As a more selective nicotinic agonists, we applied 30 µM levamisole to Fluo-3AM treated intestines to look for evidence of functional activation of L-type receptors to be indicated by the expression the subunit message described earlier. Application of levamisole produced a rise in Ca2+ amplitude, but generated another distinctive Fluo-3 response profile, which contrasted to that of 96

acetylcholine and CaCl2. Levamisole generated changes in fluorescence took a shorter time to reach peak compared to acetylcholine, averaging at 316 ± 15 s (Fig. 2.6E & F), but still slower time than 10 mM CaCl2. Additionally, unlike CaCl2 and acetylcholine, the levamisole generated responses declined immediately after reaching peak even when in the continued presence of the stimulus (Fig. 2.6E). We also quantified the average changes in Fluo-3 intensity; levamisole treatment resulted in significantly smaller increases in fluorescence, 8 ± 0.39%, than acetylcholine, 12 ± 0.41% of (Fig. 2.6G).

To see if there were differences in the areas of the intestines activated by acetylcholine and levamisole, we calculated the percentage of regions (50 x 50 µm squares) of the intestine that increased in fluorescence following the application of CaCl2, acetylcholine, and levamisole.

Fig. 2.6H, shows that 100% of the regions tested produced responses to CaCl2. Acetylcholine and levamisole also produced responses in 80% and 88% of the regions examined, respectively.

There was no significant difference suggesting receptors responding to levamisole (the L-type receptors) and other cholinergic receptors are distributed in a similar way in most of the cells of the intestine.

To verify that the observed calcium signals are a result of manipulation of nAChRs and not through other receptors such as muscarinic G proteins, we PCR screened the intestines of five adult female A. suum worms to determine if they express any G protein- linked acetylcholine receptors (GARs). A. suum is known to express a GAR protein, Asu-GAR-1 with an alternatively spliced isoform of the receptor both in the head and tail regions (Kimber et al., 2009). Our RT-

PCR results detected that the receptor is expressed in the intestine (Fig. 2.7A). As mentioned, acetylcholine is a non-selective cholinergic agonist and the observed acetylcholine signal may be partially mediated through GAR-1 as well as nAChRs. To determine if levamisole is signaling 97 solely through nAChRs, we treated intestinal samples with the nicotinic receptor antagonist mecamylamine to block nAChR mediated signaling. First, we exposed tissues to 10 µM mecamylamine and saw no changes in the calcium signal (Fig. 2.7B & D). Additionally, when

10 µM mecamylamine and 30 µM levamisole were simultaneously applied we also observed no changes in Fluo-3 fluorescence (Fig. 2.7C & D). Lastly, we applied 30 µM levamisole and upon an increase in the Ca2+ signal we co-applied 10 µM mecamylamine and saw that the signal was terminated, often to a level below the resting level as the intracellular Ca2+ is adjusted by uptake processes (Fig. 2.7E). These data demonstrate that levamisole stimulates Ca2+ signals in the intestine by signaling through nAChRs and not GARs.

2.5 Discussion

2.5.1 GAR-1s, nAChRs and a paracrine function

In this study we have observed: message of both nAChR subunits for Asu-unc-29, Asu- unc-63, Asu-unc-38, Asu-acr-8 and the G protein-linked acetylcholine receptor (GAR), Asu-gar-

1 in the cells of A. suum intestine. We have also observed that acetylcholine and levamisole produced different Ca2+ signal profiles indicating that they activate two different sets of receptors. The latency before the beginning of the levamisole calcium response, Fig. 2.6, was longer but the time to peak was shorter than that of acetylcholine. Acetylcholine is a flexible molecule and a non-selective cholinergic agonist: it can activate a range of acetylcholine receptors including G-protein coupled receptors and different types of nAChRs. Levamisole is a more rigid molecule: it is a selective L-type nAChR agonist in nematodes (Buxton et al., 2014;

Verma et al., 2017). Together, these observations indicate the presence and activation of two or more receptor signaling pathways. We interpret these observations to suggest that acetylcholine activates both G-protein acetylcholine receptors (Asu-GAR-1) and nAChRs, and that levamisole activates nAChRs that are blocked by mecamylamine. 98

Given that nematode intestine cells lack direct innervation, it is possible that intestinal

GAR-1s and nAChRs are involved in a paracrine function, responding to acetylcholine released from adjacent cells. The presence of GARs, nAChRs and acetylcholine synthesis, secretion and degradation is seen in other non-excitable cells such as: human and rat bronchial epithelial cells

(Cazzola et al., 2016; Maus et al., 1998); human keratocytes (Slonieka et al., 2015) and skin keratinocytes (Grando et al., 1993; 1995). Mellanby (1955) has detected the presence of acetylcholine in Ascaris tissues and Lee (1962) has detected cholinesterase in the intestine and secretions from of Ascaris intestinal cells. These observations support the hypothesis of a paracrine function for acetylcholine and the receptors we have observed in the intestine. Co- ordination of digestive enzyme secretion and nutrient absorption may be by signaling mediated by GAR-1s and nAChRs of the intestinal cells.

2.5.2 Different distributions of nAChRs subunit message in the basolateral and central regions of the columnar intestine cells

The nAChR subunit message for Asu-unc-29, Asu-unc-63, Asu-unc-38 and Asu-acr-8 was found in the somatic muscle and in the intestinal cells of A. suum. The protein subunits from this message produce the putative L-subtype levamisole receptor, and in different combinations produce different receptor subtypes (Buxton et al, 2014; Verma et al., 2017). Our quantitative

PCR analysis revealed that the subunit mRNA levels were differentially expressed in the muscle bag region and intestine within the intestinal columnar cells. In addition, RNAscope in situ hybridization measurements showed us that the distribution of the message for these subunits was not uniform within the columnar cells of the intestine. The message for Asu-unc-29, Asu- unc-63, Asu-unc-38 and Asu-acr-8, subunits of the putative L-type receptor, had a similar density in the region of the basolateral membrane of the columnar intestinal cells while Asu-unc-38 and

Asu-acr-8 had a much higher density than Asu-unc-29 and Asu-unc-63 in the central region of 99 the intestinal cells. The relationship between the level of mRNA expression and level of protein expressed in steady-state conditions are usually positively correlated (Liu, 2016). If we use this correlation, the different subunit message levels predict that we have combinations of nAChR subunits in the basolateral region of the intestinal cells that contrast with the combinations of the subunits in the central region of the columnar cells. These observations then suggest that there are two or more subtypes of nAChRs involved in activating the columnar intestinal cells: one subtype in the central region with Asu-unc-38 and Asu-acr-8 with less or no Asu-unc-29, Asu- unc-63 subunits; the other subtype with Asu-unc-38, Asu-acr-8, Asu-unc-29 and Asu-unc-63 subunits like the L-subtype in the basolateral region.

2.5.3 Possible function of the nAChR subtypes located in the basolateral region of the intestinal cells

Given their location, adjacent to the perienteric space, it is possible that basolateral nAChRs are involved in signaling between the muscle bags and the intestine. Acetylcholine released into the perienteric space from the bag regions of the muscle cells or from the intestinal cells can enter the perienteric canals. The perienteric canals can allow distribution of materials

(chemicals and nutrients) along the length of worm. Acetylcholine released into the perienteric space would simultaneously stimulate the L-subtype receptors present on the bag region of the

Ascaris muscle (Qian et al., 2006) and receptors present on the basolateral region of the intestinal cells. The bag region of the muscle is filled with glycogen and takes up nutrients released from the adjacent intestine requiring coordination between muscle bags and cells of the intestine.

2.5.4 Possible function of the nAChR subtypes located in the central region of the intestinal cells

The location of the nAChR subtypes in the central region of the columnar cells suggests an interaction between adjacent intestinal cells rather than with the muscle cells underneath. 100

These central nAChRs would facilitate a co-ordination of digestive enzyme secretion and nutrient absorption mediated by central region nAChRs of the columnar intestinal cells.

2.5.5 Levamisole and Cry5B

Cry5B is a Cry protein recovered from Bacillus thuringiensis that has significant anthelmintic actions. Given the widespread and very effective use of Bt Cry toxins as insecticide for protection of corn and cotton plants, it is hoped that nematocidal Cry toxins including Cry5B can be developed and used as anthelmintics. Cry5B has potent toxic effects on C. elegans and on a range of parasitic nematodes including Ascaris suum (Hu et al., 2010; Urban et al., 2013; Hu et al., 2018). The mode of action of Cry5B involves its activation by a gut protease in the nematode, binding to a nematode specific cadherin CDH-8, uptake by intestinal cells dependent on the glycolipid gene bre-5, and then a pore-forming process that damages the permeability of the intestine (Peng et al., 2018). Interestingly, nAChR agonists like levamisole and Cry5B have a synergistic anthelmintic action with levamisole increasing the potency of Cry5B (Hu et al., 2010) but a mechanism for this synergy has yet to be established.

Here we suggest that the action of levamisole on the intestine of the nematode may be to stimulate the digestion processes leading to the release of proteases and activation of Cry5B and its absorption. The activation of digestion by levamisole would then enhance the uptake of nematode intestinal contents and enhance the internalization of the activated Cry5B facilitating pore formation and the anthelmintic action of Cry5B.

We point out that benzimidazole anthelmintics also have an action on the intestinal cells of Ascaris (Borgers and De Nollin, 1975), presumably due to the disruption of microtubulins with binding to β-tubulin (Borgers et al., 1975). Uptake of the benzimidazoles by the nematode parasite is also anticipated to be increased by the cholinergic anthelmintics like levamisole;

101 levamisole is thus predicted to have an additive effect on the anthelmintic action of benzimidazoles.

2.6 Conclusion

To conclude, we have found evidence that a nematode parasite intestine expresses functioning nAChRs that respond to the cholinergic anthelmintic levamisole. These nAChRs are likely to include different subtypes, some of which may stimulate the processes of digestion and enhance the efficacy of other classes of gut acting anthelmintics.

2.7 Declaration of competing interests

The authors declare no conflict of interests.

2.8 Acknowledgments

We would like to acknowledge, NIH R01AI047194-17 and 5R21AI13967 to RJM, the E.

A. Benbrook Foundation for Pathology and Parasitology for support to RJM, NIH

R21AI092185-01A1 to APR and the College of Agriculture and Life Sciences for support for

MMcH. The funding agencies had no role in the design, execution or publication of this study.

The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infectious Diseases.

2.9 Author Contribution

M.M., P.W., S.V., J.A.P.C., A.P.R., and R.J.M. conceived the research idea and hypothesis; M.M. performed all experiments except calcium image; P.W., conducted calcium imaging; M.M analyzed and interpreted data and wrote the manuscript; P.W., S.V., J.A.P.C.,

A.P.R., and R.J.M provided assistance with data interpretation and editorial suggestions.

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2.10 Figures and Tables

Figure 2.1. Hematoxylin and Eosin-stained transverse section through Ascaris suum. The various regions are the cuticle (C); hypodermis (H); lateral lines (L); contractile spindles (S); arms (A) dorsal nerve cord (DN); ventral nerve cord (VN); muscle bags (B); Canals containing perienteric fluid (Cn) and intestine (I).

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Figure 2.2. Localization of nAChR subunits in the muscle bag region and intestine of A. suum. RT-PCR analysis of muscle bag (1b, 2b, 3b, 4b, 5b) and intestine (1i, 2i, 3i, 4i, 5i) of five separate adult female A. suum worms. Each lane represents an individual worm that muscle bags and intestinal tissue was taken for analysis. M = FastRuler Middle Range DNA Ladder (Thermo Fisher Scientific) and n.c = negative control. (A) Asu-unc-29 (B) Asu-unc-63 (C) Asu-unc-38 (D) Asu-acr-8. Note the reduced intensity of the bands (encircled white) with Asu-unc-29 (A) from the intestine. 104

Figure 2.3. Differential expression of nAChR subunits in muscle and intestine of A. suum. Bar charts (expressed as mean ± SEM ) demonstrating qPCR experiments of the relative mRNA levels of Asu-unc-38 (black bar), Asu-acr-8 (blue bar), Asu-unc-29 (red bar) and Asu-unc-63 (green bar) in: (A) muscle bag region (n = 3 individual worms) and (B) intestinal tissue (n = 3 individual worms) of A. suum. qPCR experiments were repeated three times for each gene: 3 biological replicates each with 3 technical replicates). For muscle bags, analysis revealed that Asu-unc-38 (32.83 ± 0.62, n = 3) had the highest relative mRNA levels than Asu-acr-8 (1.23 ± 0.41, n= 3), Asu-unc-29 (1.63 ± 0.22, n = 3) and Asu-unc-63 (5.77± 1.19, n = 3); *** P < 0.001. Asu-unc-63 had higher mRNA levels of expression than Asu-acr-8 and Asu-unc-29; * P < 0.05. The means for the mRNA expression of the intestinal were lower but showed similar trends with Asu-unc-38 (4.97± 1.49, n = 3) being the highest, followed by Asu-acr-8 (3.37 ± 1.34, n = 3), Asu-unc-29 (0.17 ± 0.17, n = 3) and Asu-unc-63 (2.03 ± 1.53, n = 3) but the differences did not reach statistical significance (P = 0.1250).

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Figure 2.4. Subcellular localization of nAChR subunits by RNAscope in situ hybridization in A. suum intestine. Representative images of A. suum intestine. (A) Transverse section of the intestine showing the Basolateral, Nuclear, Central and Apical regions that were observed for nAChR subunit expression. (B) Negative control probe (Bacillus DapB) where there are no pink punctate dots. (C) Asu-unc-29 probe (D) Asu-unc-63 probe (E) Asu-unc-38 probe (F) Asu-acr-8 probe. Pink punctate dots (mRNA transcript) indicate positive signal.

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Both Asu-unc-29 and Asu-unc-63 (C and D) have pink punctate dots within the basolateral region only, while Asu-unc-38 and Asu-acr-8 (E and F) have higher abundance of punctate dots distributed throughout all regions of the intestine, specifically the central region. Arrows indicate individual mRNA transcripts.

Figure 2.5. Quantitative subcellular distribution of nAChR subunit mRNA transcripts in A. suum intestine. Bar charts (mean ± SEM) showing the mean density of mRNA transcripts (punctate dots)/µm2 for Asu-unc-38, Asu-acr-8, Asu-unc-29, and Asu-unc-63 in various regions of the intestine. A) Mean density of mRNA transcripts for each nAChR subunit in all intestinal tissue visible in the cross section. * Significantly different to Asu-unc-38 (P < 0.05). 107

B) Mean density of mRNA transcripts for each nAChR subunit in basolateral region. * Significantly different to Asu-unc-38 (P < 0.05); **Significantly different to Asu-unc-38 (P < 0.01) and ***Significantly different to Asu-unc-38 (P < 0.001). C) Mean density of mRNA transcripts for each nAChR subunits in the Central region of A. suum intestine. *** Significantly different to Asu-unc-38 or Asu-acr-8 (P < 0.001). Asu-unc-38 (n = 36 image frames from 3 worms); Asu-acr-8 (n = 34 image frames from 3 worms); Asu-unc-29 (n = 39 image frames from 3 worms) and Asu-unc-63 (n = 39 image frames from 3 worms).

Figure 2.6. Acetylcholine and levamisole generate Ca2+ signals in Fluo-3AM treated Ascaris intestines. A) Micrographs of Ascaris suum intestinal section under white light (left), untreated fluorescing under blue light (center) and 5 μM Fluo-3AM treated under blue light (right), after 108

60-min incubation at 35°C with 10% Pluronic F-127. Key structures, tower/column and cells are highlighted. B) Representative trace of a 10mM CaCl2 stimulated signal. Grey box indicates application of the stimulus. C) Amplitudes of Ca2+ signals in intestines exposed to APF containing 500 µM CaCl2 (untreated) and 10mM CaCl2 (Black bar). **** Significantly different from 500 µM CaCl2 APF (P < 0.0001, t = 18.67, df = 925, unpaired t-test). 500 µM CaCl2 APF n = 238 recordings from 8 intestines from 8 individuals; 10mM CaCl2 n = 689 recordings from 12 intestine preparations from 6 individual females. D) Representative trace of a Ca2+ signal to 30 μM acetylcholine. Grey box indicates acetylcholine application. E) Representative trace of a Ca2+ signal to 30 μM levamisole. Grey box indicates levamisole application. F) Quantification of the time for the Ca2+ signal to reach peak after stimulus application for 10 mM CaCl2 (Black bar), 30 μM acetylcholine (White bar) and 30 μM levamisole (Grey bar). **** Significantly different to CaCl2 (CaCl2 vs acetylcholine P < 0.0001, t = 33.62, df = 986; CaCl2 vs levamisole P < 0.0001, t = 16.15, df = 916, unpaired t-tests). 10mM CaCl2 n = 689 recordings from 12 intestinal preparations from 6 individual females; acetylcholine n = 280 recordings 7 intestine from 4 individual worms; levamisole n = 241 recordings 5 intestinal preparations from 4 females. G) Amplitudes of Ca2+ signals in intestines in response to 30 μM acetylcholine (White bar) and 30 μM levamisole (Grey bar). **** Significantly different to acetylcholine (P < 0.0001, t = 7.182, df = 519 unpaired t-test). H) Average percentage of recording regions of the intestine with positive increases in Fluo-3 fluorescence to 10mM CaCl2 (Black bar), 30 μM acetylcholine (White bar) and 30 μM levamisole (Grey bar). N.S. = not significant (acetylcholine vs levamisole P = 0.6037, t = 0.5359, df = 10, unpaired t-test). All values are represented as means ± SEM.

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Figure 2.7. Intestine Asu-gar-1 message and mecamylamine inhibition of levamisole mediated Ca2+ signals. A) RT-PCR analysis shows the presence of Asu-gar-1 in the intestine (1i, 2i, 3i, 4i, 5i) of five separate adult female A. suum. B) Representative trace to 10 µM mecamylamine. Grey box indicates application of the stimulus. C) Representative trace for simultaneous application of 30µM levamisole (grey box) and 10µM mecamylamine (black bar). D) Quantification of 2+ maximum changes in Ca signal amplitude for untreated (Black bar), 10 µM mecamylamine (White bar) and 30 μM levamisole + 10µM mecamylamine (Grey bar). N.S. = not significant to 110 untreated (untreated vs mecamylamine P = 0.2278, t = 1.207, df = 578; untreated vs levamisole + mecamylamine P = 0.2365, t = 1.185, df = 506 unpaired t-test). Untreated n = 280 recordings from 10 intestinal preparations from 10 individual females; mecamylamine n = 300 recordings 5 intestine from 3 individual worms; levamisole + mecamylamine n = 230 recordings 4 intestinal preparations from 4 females. E) Representative trace the response to 30µM levamisole before co-application of 10 µM mecamylamine. Grey box indicates levamisole application; black represents mecamylamine. Note that mecamylamine inhibits the response to levamisole and produces a reduction in the Ca2+ fluorescence below the resting level due to a delayed uptake process that follows the rise in intracellular Ca2+. All values are represented as means ± SEM.

Supporting Information

Table S2.1. Primer sequences used for RT-PCR

Table S2.2. Primer sequences used between TM3 and TM4 for quantitative PCR

111

Figure S2.1. Visualization of positive control probe (Asu-gapdh) by RNAscope in situ hybridization in Ascaris suum intestine. Representative image of transverse section of A. suum intestine showing the positive control probe (Asu-gapdh) where pink punctate dots are clustered as a lawn within the tissue.

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Figure S2.2 Subcellular localization of nAChR subunits by RNAscope in situ hybridization in Ascaris suum muscle. Representative images of A. suum muscle. (A) Transverse section of the muscle Negative control probe (Bacillus DapB) where there are no pink punctate dots. (B) Positive control probe (Asu-gapdh) where pink punctate dots are clustered along the muscle bag membrane (C) Asu-unc-29 probe (D) Asu-unc-63 probe (E) Asu-unc-38 probe (F) Asu-acr-8 113 probe. Pink punctate dots (mRNA transcript) indicate positive signal from the hybridization of complementary mRNA. Black arrows indicate individual mRNA transcripts.

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CHAPTER 3. ALLOSTERIC MODULATION OF OESOPHAGOSTOMUM DENTATUM NICOTINIC ACETYLCHOLINE RECEPTORS BY MACROCYCLIC LACTONES

Mark McHugh a, Alan P. Robertson b, Jo Anne Powell-Coffman a, Richard J. Martin b, *

a Department of Genetics, Development, and Cell Biology, Iowa State University, Ames, Iowa

50011, USA

b Department of Biomedical Sciences, College of Veterinary Medicine, Iowa State University,

Ames, Iowa 50011, USA

*Corresponding author: [email protected]

Modified from a manuscript to be submitted to the International Journal of Parasitology Drugs

and Drug Resistance

3.1 Abstract

Parasitic nematode infections are an increasing problem that has caused substantial disease in humans and animals worldwide. Infections are controlled by use of commercially available anthelmintic drugs. However, misuse of these drugs has led to the development of resistance globally. Anthelmintics such as levamisole and pyrantel are known to target nicotinic acetylcholine receptors (nAChRs) which produces paralysis of the worm. Since anthelmintic resistance is of major concern in parasitic nematodes, an attempt to circumvent this problem is through use of combinations of two or more anthelmintics from different classes to increase treatment efficacy. Moreover, an understanding of the structural basis of the target sites, chemical structure of anthelmintics and receptor-drug interactions are critical components in resolving anthelmintic resistance. This can begin by screening pharmacological agents that 120 regulate receptor activity through allosteric modulatory sites. Allosteric modulatory sites have been discovered on nAChRs and are topographically distinct from the orthosteric sites where ligands bind. They indirectly modulate channel gating activity by evoking positive (PAM) or negative (NAM) effects on signal transduction and have been previously suggested to accommodate the avermectin, abamectin on parasitic nematodes nAChRs. Here we expressed two receptor subtypes from the pig parasite Oesophagostomum dentatum, namely Ode levamisole and Ode pyrantel/tribendimidine receptor in Xenopus oocytes. We then investigated the allosteric modulatory effects of the avermectins, abamectin and ivermectin and the milbemycin, moxidectin on both receptors. Our results showed that all three compounds acted as positive allosteric modulators (PAMs) on the Ode levamisole receptor. In contrast, the avermectins exhibited NAM activity, while moxidectin maintained its PAM activity on the Ode pyrantel/tribendimidine receptor. These findings highlight the significant role of the macrocyclic lactones in enhancing anthelmintic efficacy which can be further exploited for combination therapy.

3.2 Introduction

Neglected tropical diseases (NTDs), have negatively impacted the health and wellbeing of a substantial proportion of the world’s population. NTDs are caused mainly by parasitic nematodes, where they infect more than 1.4 billion people (including more than 500 million children). Infections thrive in impoverished areas where there is a lack of proper sanitation and hygiene, limited access to clean water, substandard healthcare, overpopulation, and people live in proximity with infective disease vectors and animals (Fewtrell et al., 2005; Collier, 2007; Hotez et al., 2009; Bartram and Cairncross, 2010; Struntz et al., 2014). Heavy infections are associated with malnutrition, impaired growth, reduced intellectual development, blindness and disfigurement (Crompton and Nesheim, 2002; Bethony et al., 2006; Hotez et al., 2008). The 121 diseases also prolong the vicious cycle of poverty and inequity because infected children are prevented from attending school thus reducing their chances of procuring good jobs, and infected adults are unable to provide a steady source of income (Bleakley, 2007; Hotez et al., 2009).

Effective vaccines are currently unavailable. Hence the global strategy to control morbidity due to NTDs is through mass drug administration (MDA) programs (WHO, 2019).

Anthelmintic drugs are generally used as a main form of control and include the benzimidazoles, macrocyclic lactones, imidazothiazoles, tetrahydropyrimidines, spiroindole and the amino- acetonitrile derivative. Although anthelmintic drugs have reduced infections in some populations, incorrect use of these drugs has led to therapeutic failures and widespread development of anthelmintic resistance (Prichard, 1994). Furthermore, the lack of financial incentive due to economic deprivation has reduced efforts by the pharmaceutical industry to undertake the development of anthelmintics for humans (Kyne et al., 2019; Nixon et al., 2020)

Combination therapy has been widely advocated, where two or more anthelmintics of different classes are combined to expand the efficacy spectrum (Leathwick et al., 2009; Geary et al., 2012; Lanusse et al., 2015). Additionally, this approach has been proven to be effective in delaying the onset of anthelmintic resistance (Leathwick et al., 2009; Keiser et al., 2012, 2013;

Smith, 2014). Cholinergic anthelmintics such as levamisole and pyrantel, are known to selectively bind to nematode nicotinic acetylcholine receptors (nAChRs), producing depolarization, spastic paralysis and removal of the nematode from its host via normal peristalsis

(Aubry et al., 1970; Aceves et al., 1970; Martin and Robertson, 2007). Other anthelmintics like the avermectins and milbemycins that belong to the macrocyclic lactone family, are known to act on nematode glutamate gated chloride channels (GluCls) by binding to allosteric sites on the channel subunits. This results in intracellular flux of Cl- ions leading to hyperpolarization of the 122 nematode neuromuscular system thus producing neuromuscular paralysis and inhibition of pharyngeal pumping and reproduction (Geary et al., 1993; Cully et al., 1994; Martin et al., 1996;

Sheriff et al., 2002). However, avermectins, namely, abamectin are not limited to GluCls, but have been shown to be an antagonist, or negative allosteric modulator of nAChRs in both Ascaris suum and Oesophagostomum dentatum (Puttachary et al., 2013; Abongwa et al., 2014).

Numerous studies have been undertaken that highlights allosteric modulation of the mammalian

α7 nAChRs (Gill et al., 2011, 2012; Gill-Thind et al., 2015; King et al., 2017; Gulsevin et al.,

2019) but information is sparse with regards to the nematode nAChRs. Therefore, investigating allosteric modulation of nematode nAChRs, can improve our understanding of the pharmacological mechanisms involved in drug combinations. Since abamectin is a selective agonist of GluCls and an allosteric modulator of nematode nAChRs, a feasible attempt to manage drug resistance would involve the combination of drugs from the macrocyclic lactone family with cholinergic agonists to increase drug efficacy.

Here we investigated the allosteric modulatory effects of three macrocyclic lactones, namely, abamectin, ivermectin and moxidectin on a levamisole selective (L-type) and pyrantel/tribendimindine selective receptor subtype (Pyr/Tbd-type) from the pig parasite,

Oesophagostomum dentatum. O. dentatum was our parasite of choice because they are easily maintained and passaged and serve as a model organism for the human species O. bifurcum which causes oesophagostomiasis. The L-type receptor is comprised of Ode-UNC-38, Ode-

UNC-29, Ode-UNC-63 and Ode-ACR-8, while the Pyr/Tbd subtype has all the subunits listed for the L-type, with the exception of ACR-8 (Buxton et al., 2014). The absence of ACR-8 results in difference in pharmacological properties and cholinergic anthelmintic selectivity between the receptors. Abamectin and ivermectin are both structurally similar, while moxidectin is 123 structurally dissimilar. In this regard, we developed two hypotheses that guided our experiments.

These are as follows: 1) Differences in subunit combination and stoichiometry of the receptor subtypes will impact allosteric modulation by the macrocyclic lactones; 2) Both avermectins

(abamectin and ivermectin) will have different modulatory effects on the O. dentatum nAChR receptor subtypes, when compared to the milbemycin, moxidectin due to structural similarities between the avermectins. We report that both avermectins (abamectin and ivermectin) acted as a positive allosteric modulator (PAM) on the Ode levamisole receptor, while acting as negative allosteric modulators (NAM) on the Ode-Pry/Tbd receptor subtype. However, the nature of the

PAM and NAM effect was different between both compounds. In contrast, moxidectin acted as a

PAM on both receptor subtypes. These findings are significant and improve our understanding on the prospective use of drug combination in combating anthelmintic resistance.

3.3 Materials and Methods

3.3.1 Molecular Biology

Cloning was conducted for O. dentatum (Ode-unc-38, Ode-unc-29, Ode-unc-63 and Ode- acr-8) nAChR subunits and Haemonchus contortus ancillary factors (Hco-unc-50, Hco-unc-74 and Hco-ric-3) as previously described (Abongwa et al., 2016; Buxton et al., 2014).

3.3.2 Heterologous expression of Ode-levamisole receptors and Ode-pyrantel receptors in Xenopus laevis oocytes

Defolliculated Xenopus laevis oocytes were purchased from Ecocyte Bioscience (Austin,

TX, USA). Heterologous expression of the Ode-levamisole receptor was achieved by co- injecting 1.8 ng of cRNA for each subunit (Ode-unc-38, Ode-unc-29, Ode-unc-63, Ode-acr-8) with 1.8 ng of each ancillary factor (Hco-ric-3, Hco-unc-50 and Hco-unc-74) in a total volume of 36 nL in nuclease-free water. Each oocyte was microinjected into the cytoplasm of the animal pole region using a Drummond Nanoject II microinjector (Drummond Scientific, Broomall, PA, 124

USA). After injection, oocytes were incubated at 19°C in a sterile 96-well culture plate

. containing 200 µl of incubation solution (100 mM NaCl, 2 mM KCl, 1.8 mM CaCl2 2H2O, 1 mM

. MgCl2 6H2O, 5 mM HEPES, 2.5 mM Na pyruvate, 100 U/mL penicillin and 100 µg/mL streptomycin, pH 7.5) in each well. Incubation solution was changed daily during the period of incubation. The same procedure was also conducted for the Ode-pyrantel receptor with the following subunits (Ode-unc-38, Ode-unc-29 and Ode-unc-63) and ancillary factors (Hco-ric-3,

Hco-unc-50 and Hco-unc-74). Experiments were performed on oocytes that expressed the Ode- levamisole nAChRs 5 days post injection, while oocytes expressing Ode-pyrantel nAChRs were used 7 days post injection.

3.3.4 Two-microelectrode voltage camp (TEVC) electrophysiology

Oocytes were treated with 100 µM BAPTA-AM for approximately 3 hours prior to electrophysiology recordings. This was done to prevent activation of endogenous calcium- activated chloride currents during the recording. TEVC was conducted by impaling oocytes with two microelectrodes; a current injecting electrode, Im, used to inject the required current for holding the membrane at a set voltage, and a voltage sensing electrode, Vm. The microelectrodes were pulled using a Flaming/ Brown horizontal electrode puller ((Model P-97; Sutter

Instruments, Novato, CA, USA) and filled with 3 M KCl. Each electrode tip was broken with a piece of kimwipe paper (Kimtech ScienceTM, Fisher) to achieve a resistance of 2 – 5 MΏ in

. recording solution (100 mM NaCl, 2.5 mM KCl, 1 mM CaCl2 2H2O and 5 mM HEPES, pH 7.3).

Oocytes were voltage clamped at -60 mV with an Axoclamp 2B amplifier (Molecular Devices,

Sunnyvale, CA, USA). Amplified signals were converted from analog to digital format by a

Digidata 1322A digitizer (Molecular Devices, CA, USA) and all data were acquired on a desktop computer with the Clampex 10.3 data acquisition software (Molecular Devices, Sunnyvale, CA,

USA). 125

3.3.5 Drugs

Acetylcholine chloride (ACh) and moxidectin (Mox) were purchased from Sigma-

Aldrich (St. Louis, MO, USA). Abamectin and ivermectin were purchased from Alfa Aesar

(Ward Hill, MA, USA) and MP Biomedicals, LLC (Solon, OH, USA) respectively. Fresh ACh stock solutions were prepared in recording solution daily, prior to experimentation. Stock solutions of abamectin, moxidectin and ivermectin were made in DMSO, then diluted in recording solution such that the final DMSO concentration did not exceed 0.1%.

3.3.6 Drug application

All agonist concentration-response experiments were conducted by initially applying 100 µM of acetylcholine, followed by application of increasing concentrations (0.3 – 100 µM) of acetylcholine. Each concentration was applied for 10 s and allowed to wash off for 3 mins between each application. For allosteric modulator experiments, the effects of abamectin, moxidectin and ivermectin were tested on the acetylcholine concentration-relationship in separate experiments. Briefly, an initial 100 µM acetylcholine was applied for 10 s to each oocyte, followed by a 10 min challenge with abamectin and 10 s applications of increasing concentrations of the agonist in the continued presence of abamectin. A 3 min wash off time was allowed between each application. The same procedure was repeated for moxidectin and ivermectin.

3.3.7 Data analysis

Peak current responses for each acetylcholine concentration in the absence or presence of abamectin, moxidectin or ivermectin were measured and normalized to the initial control 100

µM acetylcholine responses (for each oocyte) using Clampfit 10.3 (Molecular Devices,

Sunnyvale, CA, USA). Concentration-response relationships were analyzed by fitting log concentration-response data points with the Hill equation as previously described: where EC50 is 126

the concentration producing the half-maximum response; Rmax is the maximum response % relative to the control 100 µM ACh response and nH is the slope factor or Hill coefficient (Boulin et al., 2008). Statistical differences among treatment groups were tested using one-way analysis of variance (ANOVA), P < 0.05, followed by the Tukey multiple comparison post-hoc test. This was done using the GraphPad Prism 5.0 software (GraphPad Software, Inc., USA), and results were expressed as mean ± S.E.M.

3.4 Results

3.4.1 Effects of abamectin on Ode levamisole receptors

We tested the effects of different abamectin concentrations (0.1 µM, 1 µM and 10 µM) on Ode levamisole responses to acetylcholine. For control experiments, each acetylcholine concentration from 0.3 to 100 µM was applied for 10s, Fig 3.2A. To determine the effects of abamectin, 100 µM acetylcholine was first applied for 10s as control, followed by a 3 min wash, then perfusion with abamectin for 10 mins, after which acetylcholine applications were repeated in the presence of abamectin, Fig 3.2B. A 3 min wash period was applied between drug applications for both control and test experiments. We observed that abamectin failed to activate the Ode levamisole receptor when applied on its own, thus eliminating agonist activity on the receptor, Fig 3.2B. However, when we co-applied acetylcholine with 0.1 µM or 1 µM abamectin, we saw potentiation of Ode-levamisole responses to acetylcholine. In marked contrast, increasing the concentration of abamectin to 10 µM caused no significant change in acetylcholine responses. Fig 3.2C shows concentration-response plots for these experiments.

Overall, abamectin concentrations of 0.1 µM and 1 µM caused a significant increase in Rmax (Fig

3.2D), but did not change the EC50, whereas abamectin at 10 µM caused no change in Rmax or

EC50. Rmax and EC50 values are shown in Table 3.1. These findings suggest that abamectin is a 127 positive allosteric modulator (PAM) of the Ode levamisole receptor at low concentrations but abolishes PAM activity at higher concentrations.

3.4.2 Effects of ivermectin on Ode levamisole receptors

To test the hypothesis that ivermectin will have similar modulatory effects as abamectin on the Ode-levamisole receptor due to structural similarities between the compounds (Fig 3.1A

& B), we used the same experimental protocol previously described for abamectin. Application of ivermectin on its own did not cause activation of the receptor, thus demonstrating no agonist activity, Fig 3.3A. However, co-application of acetylcholine with ivermectin, increased the potency of acetylcholine which was evident by a significant decrease in EC50 for all the concentrations of abamectin tested, namely 0.1µM, 1µM and 10µM, Fig 3.3B. Moreover, Rmax values remained unchanged. Rmax and EC50 values are shown in Table 3.2. These results confirm that ivermectin and abamectin are both positive allosteric modulators of the Ode-levamisole receptor. However, the nature of the PAM effect was different, where ivermectin increased the potency of acetylcholine by causing a left shift in EC50, whereas abamectin improved the efficacy of acetylcholine at low concentrations by increasing the maximum response (Rmax).

3.4.3 Effects of moxidectin on Ode levamisole receptors

Since moxidectin is structurally different when compared to the avermectins (ivermectin and abamectin) (Fig 3.1C), we hypothesized that the mode of action of moxidectin would be different from the avermectins on the Ode-levamisole receptor. Hence, the effects of moxidectin on the Ode-levamisole receptor was tested using the same protocol as previously mentioned for ivermectin and abamectin. A representative trace of the experiment can be seen in Fig 3.4A. We observed that application of moxidectin on its own, failed to elicit an agonist response, Fig 3.4A.

This was similar to that previously seen for the avermectins, namely, abamectin and ivermectin.

However, co-application of acetylcholine with 0.1 µM or 10 µM moxidectin potentiated the Ode 128 levamisole responses to acetylcholine, Fig 3.4B. On the other hand, co-application of acetylcholine with 1 µM moxidectin, did not result in any change on the acetylcholine responses,

Fig 3.4B. Overall, moxidectin concentrations of 0.1 µM and 10 µM caused a significant increase in Rmax, but no change the EC50, whereas abamectin at 1 µM caused no change in Rmax or EC50.

Rmax and EC50 values are shown in Table 3.3. This suggest that moxidectin is a positive allosteric modulator of the Ode-levamisole receptor at low and high concentrations. Additionally, the nature of the PAM effect was also similar to that of abamectin, where both compounds increased the efficacy of acetylcholine by increasing the maximum response (Rmax) at a concentration of

0.1µM. Nevertheless, abamectin lacked PAM activity at higher concentrations, when compared to moxidectin.

3.4.4 Effects of abamectin on Ode pyrantel/tribendimidine receptors

Previous studies have shown that the Ode-pyrantel/tribendimidine (Ode-Pyr/Tbd) receptor is comprised of Ode-UNC-38, Ode-UNC-29 and Ode-UNC-63 nAChR subunits along with the 3 ancillary factors, Hco-RIC-3, Hco-UNC-74 and Hco-UNC-50 (Buxton et al., 2014).

The exclusion of Ode-ACR-8 made this receptor subtype more selective for pyrantel and tribendimidine as opposed to levamisole. (Buxton et al., 2014). Here we tested the effects of abamectin at concentrations of 0.1 µM, 1 µM and 10 µM on the Ode-Pyr/Tbd receptor subtype.

We used the same experimental protocol previously described for Ode levamisole receptors. A representative trace of the electrophysiology recording is shown in Fig 3.5A. Abamectin applied on its own did not activate the receptor, demonstrating no agonist activity, Fig 3.5B. Co- application of acetylcholine with abamectin at concentrations of 1 µM or 10 µM reduced the potency of acetylcholine, by causing an increase in EC50 for both concentrations, Fig 3.5C. In contrast, this inhibitory effect was not seen with 0.1 µM abamectin. The maximum response

(Rmax) remained unchanged for all the concentrations of abamectin tested, Fig 3.5D. Rmax and 129

EC50 values are shown in Table 3.4. These findings suggest that abamectin is a negative allosteric modulator (NAM) of the Ode-Pyr/Tbd receptor at concentrations that are ≥ 1µM, while lacking NAM activity at concentrations that are ≤ 0.1 µM.

3.4.5 Effects of ivermectin on Ode pyrantel/tribendimidine receptors

For these experiments, our hypothesis remained the same as previously mentioned with the Ode levamisole receptor. We hypothesized that structural similarities between ivermectin and abamectin, would yield similar allosteric modulatory effects on the Ode-Pyr/Tbd receptor. We conducted these experiments using the same protocol as previously described for the Ode levamisole receptor. Figure 3.6A shows a representative trace of our results. We observed that application of ivermectin on its own did not evoke agonist action on the receptor, Fig 3.6A.

Moreover, when co-applied with acetylcholine, 0.1 µM ivermectin caused inhibition in acetylcholine responses, where there was significant reduction in the maximum response (Rmax) and no change in EC50. In contrast 1 µM and 10 µM ivermectin had no effect on acetylcholine responses, Fig 3.6B. Rmax and EC50 values are shown in table 3.5. Taken together, these results indicate that ivermectin at 0.1 µM is acting as a negative allosteric modulator, by way of non- competitive antagonism on the Ode-Pry/Tbd receptor. On the other hand, allosteric modulation is not exhibited by ivermectin at concentrations of 1 µM and 10 µM.

3.4.6 Effects of moxidectin on Ode pyrantel/tribendimidine receptors

To determine the effects of moxidectin on expressed Ode-Pry/Tbd receptors, we used the same experimental protocol described for Ode levamisole receptors. Moxidectin alone did not activate the Ode-Pyr/Tbd receptor, thus ruling out agonist action, Fig 3.7A. Co-application of acetylcholine with 10 µM moxidectin potentiated acetylcholine responses, which lead to a significant increase in the maximum response (Rmax), but no change in EC50, (Fig 3.7B & C).

Rmax and EC50 values are shown in table 3.6. It was also observed that concentrations of 0.1 µM 130 and 1 µM moxidectin had no effect on acetylcholine responses. These results highlight moxidectin as a positive allosteric modulator of the Ode-Pyr/Tbd receptor, which is different from the NAM effects seen with abamectin and ivermectin.

3.5 Discussion

3.5.1 Diversity of parasitic nematodes nAChR subtypes

In this study, we expressed two receptor subtypes from O. dentatum in Xenopus laevis oocytes that differ in subunit combination and cholinergic selectivity. This included the Ode- levamisole (Ode-Lev) receptor subtype that is comprised of Ode-UNC-38, Ode-UNC-29, Ode-

UNC-63 and Ode-ACR-8 subunits which is selective to levamisole, and the Ode-

Pyrantel/tribendimidine (Ode-Pyr/Tbd) subtype that consists of Ode-UNC-38, Ode-UNC-29 and

Ode-UNC-63 subunits which is selective to pyrantel and tribendimidine. The main goal of this experiment was to determine the allosteric modulatory effects of the macrocyclic lactones, namely, abamectin, ivermectin and moxidectin on both receptor subtypes.

Macrocyclic lactones are known to target glutamate-chloride channels (GluCls) in an irreversible manner, leading to flaccid paralysis of neuromuscular systems (Geary et al., 1993;

Martin et al., 1996, 1997; Sheriff et al., 2002). However, studies conducted with abamectin have shown that the macrocyclic lactone is not limited to GluCls but also interact with nAChRs causing allosteric modulation of the receptor (Puttachary et al., 2013; Abongwa et al., 2014).

With this in mind, we believe that other compounds in the macrocyclic lactone family also possess the capacity to modulate various nAChR receptor subtypes. We report that both the avermectins, namely, ivermectin and abamectin and the milbemycin, moxidectin displayed allosteric modulatory effects on the Ode-lev and Ode-Pyr/Tbd receptor subtypes.

131

3.5.2 The avermectins (abamectin and ivermectin) and the milbeymycin, moxidectin are positive allosteric modulators (PAMs) of the Ode levamisole receptor

We found that the avermectins, namely, abamectin and ivermectin, and the milbemycin, moxidectin were positive allosteric modulators of the Ode-lev receptor. Interestingly, the nature of the PAM activity was similar for both abamectin and moxidectin, but different for ivermectin.

Abamectin and moxidectin increased the efficacy of acetylcholine (increase in Rmax), whereas ivermectin increased the potency of acetylcholine (left shift in EC50). This difference in PAM activity could possibly be attributed to minor structural differences between abamectin and ivermectin, since moxidectin exhibited similar characteristics as abamectin.

Abamectin is composed of a mixture of avermectin B1a (> 90%) and avermectin B1b (<

10%), whereas ivermectin is a chemically reduced 22, 23-dihydro derivative of abamectin, and is a mixture of 22, 23-dihydroavermectin B1a (> 90%) and 22, 23-dihydroavermectin B1b (< 10%)

(Campbell, 1989). Both molecules display structural similarities but have minor differences only at position C22 and C23 where hydrogen is absent in abamectin due to the presence of an olefinic bond (unsaturated hydrocarbon), while hydrogen is present in ivermectin due to hydration of the double bonds (Campbell et al., 1989). Therefore, it can be suggested that the presence of hydrogen in ivermectin, results in slightly different interactions with the Ode-lev receptor, thus resulting in a reduction in EC50 rather than potentiation of acetylcholine responses as seen with abamectin and moxidectin.

3.5.3 The avermectins (abamectin and ivermectin) are negative allosteric modulators (NAMs) of the Ode-Pyr/Tbd receptor, while moxidectin exhibits positive allosteric modulatory effects (PAM).

The Ode-Pyr/Tbd receptor is comprised of similar subunits as the Ode-lev receptor subtype, with the exception of ACR-8 being absent from the Ode-Pyr/Tbd (Ode-38-29-63) subtype and present in the Ode-lev (Ode-38-29-63-8) receptor. This difference in receptor 132 subunit combination and stoichiometry, results in differences in receptor pharmacology and selectivity. We observed that both abamectin and ivermectin antagonized the Ode-Pyr/Tbd receptor subtype through negative allosteric modulation (NAM), while moxidectin maintained its

PAM activity as previously seen with the Ode-lev receptor. It was also noteworthy that the nature of the NAM effect was slightly different between the avermectins, whereas moxidectin’s

PAM effect was consistent (increase in Rmax).

We believe that the presence of the disaccharide group on the avermectins, and the absence of the ACR-8 subunit from this receptor subtype, resulted in slight differences in the interaction of the molecules with the receptor, thus reversing the PAM effect previously seen with the Ode-lev receptor. Furthermore, the presence of hydrogen at position C22 and C23 on ivermectin may have also caused differences in the nature of the NAM effect where ivermectin reduced the efficacy of acetylcholine (reduction in Rmax) while abamectin reduced acetylcholine potency (right shift in EC50). Moxidectin on the other hand maintained its PAM effect because it is unaffected by the absence of ACR-8. Additionally, the absence of the disaccharide substituent and the presence of the methoxime on the spiroketal ring of moxidectin may have created unique interactions with this receptor subtype which may be different from that seen in GluCls. Hence, these differences may have allowed moxidectin to bind to a PAM site rather than a NAM site.

3.5.4 Use of drug combinations in combating anthelmintic resistance in parasitic nematodes

Drug combinations of different anthelmintic classes have been advocated as a suitable approach in delaying the onset of drug resistance in parasitic nematodes. Significant interest has also been shown for compounds that have allosteric modulatory activity, which facilitates the use of lower drug doses, thus reducing toxicity, minimizing the development of resistance, and improving drug efficacy. Hence, the use of pharmacology-based information is pivotal in the 133 design and utilization of successful strategies to combat anthelmintic resistance. The avermectins and milbemycins are widely used in the control of parasitic nematodes in humans, livestock and companion animals (Prichard and Geary, 2019). Moreover, resistance to these drugs is of increasing concern. Hence, our results reveal that combination of the macrocyclic lactones with cholinergic agonists such as levamisole or pyrantel would yield similar effects as when the natural agonist, acetylcholine is present, thus improving drug efficacy in vivo. However, detailed interpretation of these results is somewhat limited due to the lack of high-resolution crystal structures or homology models of these receptor subtypes that include the pore-forming transmembrane domains. As a consequence, we are currently unable to predict the precise interactions that occur for abamectin, ivermectin and moxidectin in the presence of the agonist, acetylcholine, levamisole or pyrantel. Nonetheless, studies of the mammalian α7 receptor has shown that compounds differing only in methyl substitution of a single aromatic ring exhibited five distinct pharmacological effects (type I PAMs, type II PAMs, NAMs, allosteric agonists and

SAMs) on the receptor (Gill-Thind et al., 2014). Taken together, structural differences between each compound, nAChR subunit combination and stoichiometry are critical factors to consider when evaluating the utilization of drug combinations.

3.6 Conclusion

We conclude that the avermectins (abamectin and ivermectin) and the milbemycin, moxidectin are allosteric modulators of Ode-Lev and Ode-Pyr/Tbd receptors. It was noteworthy that moxidectin maintained its PAM activity across receptor subtypes, while abamectin and ivermectin were PAMs on the Ode-Lev receptor and NAMs on the Ode-Pyr/Tbd receptor subtype. These findings suggest that the Ode-Lev receptor possess only PAM sites for both the avermectins and milbemycin, whereas the Ode-Pyr/Tbd subtype has one PAM site for 134 moxidectin and multiple NAM sites for abamectin and ivermectin. These allosteric modulation sites can be further exploited for anthelmintic therapy.

3.7 Declaration of competing interest

The authors declare no conflict of interests

3.8 Acknowledgement

We acknowledge, the NIH National Institute of Allergy and Infectious Diseases grants

R01AI047194-17 and 5R21AI13967 to RJM, the E. A. Benbrook Foundation for Pathology and

Parasitology for support to RJM, NIH R21AI092185-01A1 to APR, the College of Agriculture and Life Sciences and the College of Liberal Arts and Sciences for support for MMcH.

3.9 Author Contribution

M.M., A.P.R., J.A.P.C., and R.J.M. conceived the research idea and hypothesis; M.M. performed research, analyzed and interpreted data and wrote the manuscript; A.P.R., J.A.P.C and

R.J.M. provided assistance with data interpretation and editorial suggestions. 135

3.10 Figures and Tables

Figure 3.1. Chemical structure of avermectins and milbemycin anthelmintics. A: Structure of abamectin. B: Structure of ivermectin. C: Structure of moxidectin.

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Figure 3.2. Acetylcholine concentration-response relationships in the presence of 0.1 µM, 1 µM and 10 µM abamectin. A: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63:Ode-ACR-8 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by 10 seconds application of different acetylcholine concentrations, from 0.3 µM to 100 µM. B: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode- UNC-63:Ode-ACR-8 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by a 10 minute application of abamectin (0.1 µM) and finally a 10 seconds application of different acetylcholine concentrations in the continued presence of abamectin. C: Concentration-response plots of acetylcholine in the absence (n = 6, black) and presence of 0.1 µM abamectin (n = 6, blue), 1 µM abamectin (n = 6, red) and 10 µM abamectin (n = 6, green). Results were normalized to 100 µM acetylcholine current responses and 137 expressed as mean ± S.E.M. Note that 0.1 µM and 1 µM abamectin caused potentiation of current responses to acetylcholine. D: Bar chart showing the mean ± S.E.M. of the maximum current responses (Rmax) for acetylcholine and the different abamectin concentrations. Rmax for 0.1 µM abamectin (***P < 0.001, n = 6, blue) and 1 µM abamectin (**P < 0.01, n = 6, red) were significantly higher than Rmax for acetylcholine alone (n = 6, black). There was no significant difference in Rmax between 10 µM abamectin (n = 6, green) and acetylcholine alone (n = 6, black).

Figure 3.3. Acetylcholine concentration-response relationships in the presence of 0.1 µM, 1 µM and 10 µM ivermectin. A: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63:Ode-ACR-8 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by a 10 minute application of ivermectin (0.1 µM) and finally a 10 seconds application of different acetylcholine 138 concentrations in the continued presence of ivermectin. B: Concentration-response plots of acetylcholine in the absence (n = 6, black) and presence of 0.1 µM ivermectin (n = 6, green), 1 µM ivermectin (n = 6, red) and 10 µM ivermectin (n = 6, orange). Results were normalized to 100 µM acetylcholine current responses and expressed as mean ± S.E.M. Note that all concentrations, namely 0.1 µM, 1 µM and 10 µM ivermectin caused a left shift or reduction in EC50 of current responses to acetylcholine. C: Bar chart showing the mean ± S.E.M. of the maximum current responses (Rmax) for acetylcholine and the different ivermectin concentrations. Rmax for 0.1 µM ivermectin (n = 6, green), 1 µM ivermectin (n = 6, red) and 10 µM ivermectin (n = 6, orange) were not significantly different than Rmax for acetylcholine alone (n = 6, black).

Figure 3.4. Acetylcholine concentration-response relationships in the presence of 0.1 µM, 1 µM and 10 µM moxidectin. A: Representative trace (inward currents, holding potential -60 mV, 139 from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63:Ode-ACR-8 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by a 10 minute application of moxidectin (10 µM) and finally a 10 seconds application of different acetylcholine concentrations in the continued presence of moxidectin. B: Concentration-response plots of acetylcholine in the absence (n = 6, black) and presence of 0.1 µM moxidectin (n = 6, brown), 1 µM moxidectin (n = 6, pink) and 10 µM moxidectin (n = 6, bright blue). Results were normalized to 100 µM acetylcholine current responses and expressed as mean ± S.E.M. Note that 0.1 µM and 10 µM moxidectin caused potentiation of current responses to acetylcholine. C: Bar chart showing the mean ± S.E.M. of the maximum current responses (Rmax) for acetylcholine and the different moxidectin concentrations. Rmax for 0.1 µM moxidectin (**P < 0.01, n = 6, brown) and 10 µM moxidectin (**P < 0.01, n = 6, bright blue) were significantly higher than Rmax for acetylcholine alone (n = 6, black). There was no significant difference in Rmax between 1 µM moxidectin (n = 6, pink) and acetylcholine alone (n = 6, black).

140

Figure 3.5. Acetylcholine concentration-response relationships in the presence of 0.1 µM, 1 µM and 10 µM abamectin. A: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by 10 seconds application of different acetylcholine concentrations, from 0.3 µM to 100 µM. B: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by a 10 minute application of abamectin (10 µM) and finally a 10 seconds application of different acetylcholine concentrations in the continued presence of abamectin. C: Concentration-response plots of acetylcholine in the absence (n = 6, black) and presence of 10 µM abamectin (n = 5, light blue). Plots for 1 µM and 10 µM abamectin were left out to prevent overcrowding of the figure. Results were normalized to 100 µM acetylcholine current responses and expressed as mean ± 141

S.E.M. Note that 1 µM (not shown) and 10 µM abamectin caused inhibition of current responses to acetylcholine. D: Bar chart showing the mean ± S.E.M. of the maximum current responses (Rmax) for acetylcholine and the different abamectin concentrations. There was no significant difference in Rmax for 0.1 µM abamectin (n = 6, bright green), 1 µM abamectin (n = 6, red) and 10 µM abamectin (n = 5, light blue) when compared to the Rmax for acetylcholine alone (n = 6, black).

Figure 3.6. Acetylcholine concentration-response relationships in the presence of 0.1 µM, 1 µM and 10 µM ivermectin. A: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by a 10 minute application of ivermectin (0.1 µM) and finally a 10 seconds application of different acetylcholine 142 concentrations in the continued presence of ivermectin. B: Concentration-response plots of acetylcholine in the absence (n = 6, black) and presence of 0.1 µM ivermectin (n = 6, green), 1 µM ivermectin (n = 6, pink) and 10 µM ivermectin (n = 6, blue). Results were normalized to 100 µM acetylcholine current responses and expressed as mean ± S.E.M. Note that 0.1 µM ivermectin caused inhibition of current response to acetylcholine, whereas 1 µM and 10 µM ivermectin had no significant effect on acetylcholine current responses. C: Bar chart showing the mean ± S.E.M. of the maximum current responses (Rmax) for acetylcholine and the different ivermectin concentrations. Rmax for 0.1 µM ivermectin (**P < 0.01, n = 6, green) was significantly lower than Rmax for acetylcholine alone (n = 6, black). In contrast, Rmax for 1 µM ivermectin (n = 6, pink) and 10 µM ivermectin (n=6, blue) were not significantly different than Rmax for acetylcholine alone (n = 6, black).

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Figure 3.7. Acetylcholine concentration-response relationships in the presence of 0.1 µM, 1 µM and 10 µM moxidectin. A: Representative trace (inward currents, holding potential -60 mV, from oocytes expressing Ode-UNC-38:Ode-UNC-29:Ode-UNC-63 subunits) of the 10 seconds application of the control 100 µM acetylcholine, followed by a 10 minute application of moxidectin (10 µM) and finally a 10 seconds application of different acetylcholine concentrations in the continued presence of moxidectin. B: Concentration-response plots of acetylcholine in the absence (n = 6, black) and presence of 0.1 µM moxidectin (n = 6, dark yellow), 1 µM moxidectin (n = 6, light blue) and 10 µM moxidectin (n = 6, red). Results were normalized to 100 µM acetylcholine current responses and expressed as mean ± S.E.M. Notice that 10 µM moxidectin caused potentiation of current responses to acetylcholine. C: Bar chart showing the mean ± S.E.M. of the maximum current responses (Rmax) for acetylcholine and the different moxidectin concentrations. Rmax for 10 µM moxidectin (*P < 0.05, n = 5, red) was significantly higher than Rmax for acetylcholine alone (n = 6, black). There was no significant difference in Rmax for 0.1 µM moxidectin (n = 6, dark yellow) and 1 µM moxidectin (n = 6, light blue) when compared to acetylcholine alone (n = 6, black).

Table 3.1. EC50, Rmax, nH and n numbers for acetylcholine in the absence and presence of abamectin on Ode-UNC-38:Ode-UNC-29:Ode-UNC-63:Ode-ACR-8

Drug EC50 Rmax nH n (mean ± S.E.M, μM) (mean ± S.E.M, % 100 μM acetylcholine response) Acetylcholine 2.9 ± 0.2 104.2 ± 4.27 1.0 ± 0.1 6 + 0.1 μM abamectin 3.4 ± 0.2 127.1 ± 3.5 1.0 ± 0.1 6 + 1 μM abamectin 3.5 ± 0.1 121.5 ± 2.1 0.9 ± 0.1 6 + 10 µM abamectin 2.5 ± 0.2 106.4 ± 1.8 1.0 ± 0.0 6

Table 3.2. EC50, Rmax, nH and n numbers for acetylcholine in the absence and presence of ivermectin on Ode-UNC-38:Ode-UNC-29:Ode-UNC-63:Ode-ACR-8

Drug EC50 Rmax nH n (mean ± S.E.M, (mean ± S.E.M, % 100 μM μM) acetylcholine response) Acetylcholine 2.9 ± 0.2 104.2 ± 4.2 1.0 ± 0.0 6 + 0.1 μM ivermectin 2.1 ± 0.1 113.1 ± 1.6 1.0 ± 0.0 6 + 1 μM ivermectin 1.7 ± 0.0 104.3 ± 1.5 1.0 ± 0.1 6 + 10 µM ivermectin 1.4 ± 0.1 109.9 ± 1.4 1.0 ± 0.0 6

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Table 3.3. EC50, Rmax, nH and n numbers for acetylcholine in the absence and presence of moxidectin on Ode-UNC-38:Ode-UNC-29:Ode-UNC-63:Ode-ACR-8

Drug EC50 Rmax nH n (mean ± S.E.M, μM) (mean ± S.E.M, % 100 μM acetylcholine response) Acetylcholine 2.9 ± 0.2 104.2 ± 4.2 1.0 ± 0.0 6 + 0.1 μM moxidectin 2.8 ± 0.4 131.6 ± 5.3 0.9 ± 0.0 6 + 1 μM moxidectin 1.9 ± 0.1 107.7 ± 3.3 1.0 ± 0.0 6 + 10 µM moxidectin 2.2 ± 0.3 131.3 ± 4.3 1.0 ± 0.0 6

Table 3.4. EC50, Rmax, nH and n numbers for acetylcholine in the absence and presence of abamectin on Ode-UNC-38:Ode-UNC-29:Ode-UNC-63

Drug EC50 Rmax nH n (mean ± S.E.M, μM) (mean ± S.E.M, % 100 μM acetylcholine response) Acetylcholine 13.1 ± 1.3 110 ± 4.3 1.0 ± 0.0 6 + 0.1 μM abamectin 16.4 ± 0.8 127 ± 3.7 1.0 ± 0.0 6 + 1 μM abamectin 20.0 ± 1.9 123 ± 6.0 0.7 ± 0.1 6 + 10 µM abamectin 20.5 ± 2.0 122 ± 8.7 0.8 ± 0.0 5

Table 3.5. EC50, Rmax, nH and n numbers for acetylcholine in the absence and presence of ivermectin on Ode-UNC-38:Ode-UNC-29:Ode-UNC-63

Drug EC50 Rmax nH n (mean ± S.E.M, μM) (mean ± S.E.M, % 100 μM acetylcholine response) Acetylcholine 13.1 ± 1.3 110 ± 4.3 1.0 ± 0.0 6 + 0.1 μM ivermectin 10.8 ± 1.0 85 ± 5.5 1.0 ± 0.0 6 + 1 μM ivermectin 15.7 ± 0.8 103 ± 3.8 1.0 ± 0.1 6 + 10 µM ivermectin 16.6 ± 1.8 127 ± 5.2 1.0 ± 0.0 6

Table 3.6. EC50, Rmax, nH and n numbers for acetylcholine in the absence and presence of moxidectin on Ode-UNC-38:Ode-UNC-29:Ode-UNC-63

Drug EC50 Rmax nH n (mean ± S.E.M, μM) (mean ± S.E.M, % 100 μM acetylcholine response) Acetylcholine 13.1 ± 1.3 110 ± 4.3 1.0 ± 0.0 6 + 0.1 μM moxidectin 13.7 ± 0.5 99 ± 4.3 1.1 ± 0.0 6 + 1 μM moxidectin 15.7 ± 0.7 118 ± 4.1 1.0 ± 0.0 6 + 10 µM moxidectin 14.5 ± 1.8 132 ± 6.6 1.0 ± 0.0 5

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CHAPTER 4. ANTHELMINTIC RESISTANCE AND HOMEOSTATIC PLASTICITY: BRUGIA MALAYI

Sudhanva S. Kashyap1, Saurabh Verma1, Mark McHugh2, Mengisteab Wolday1, Paul

Williams1, Alan P. Robertson1 and Richard J. Martin1*

1Department of Biomedical Sciences, Iowa State University, Ames, IA 50011, USA

2Department of Genetics, Development and Cell Biology, Iowa State University, Ames,

IA 50011, USA

Modified from a manuscript under review in PLOS Biology

4.1 Abstract

Homeostatic plasticity refers to the capacity of excitable cells to regulate their activity to make compensatory adjustments to long-lasting stimulation. It is found across the spectrum of vertebrate and invertebrate species including nematodes and is driven by changes in cytosolic calcium; it has not been explored in parasitic nematodes when exposed to anthelmintic drugs.

The speed and level of habituation and adaptation to anthelmintics will bear on their ability to resist treatment. Here we have studied the adaptation of Brugia malayi to exposure to levamisole that activates muscle AChR ion-channels. We found that there were three phases of the Brugia malayi motility response as they adapt to levamisole: i) an initial spastic paralysis, lasting up to

20 minutes; ii) a flaccid paralysis that follows lasting up to 180 minutes and finally; iii) a recovery of motility with loss of sensitivity to levamisole by 4 hours. In calcium fluorescence experiments using fluo-3 and patch-clamp, we saw that levamisole activated AChR ion-channels and a modest increase sarcolemma calcium that was then followed by a large increase in calcium. Patch-clamp experiments after recovery of motility at 4 hours revealed that muscle 150 responses to levamisole and other cholinergic anthelmintics had desensitized. Molecular analysis of the 4-hour worms revealed an increase in the expression of unc-38 AChR subunits but no other muscle AChR subunits, while nra-2 that codes for an ER retention protein expression was decreased. RNAi of nra-2 produced: i) a significant reduction of the levamisole currents; ii) no significant change in the peak acetylcholine currents and; iii) a quicker recovery of motility in levamisole suggesting that loss of NRA-2 selectively favors the insertion of levamisole insensitive AChRs (and loss of L-subtypes) in muscle. Surprisingly, knockdown of single AChR subunits did not affect the spastic and flaccid paralysis responses to levamisole but unc-38, acr-16 and acr-26 were required for recovery of motility and habituation. These experiments show that: (i) the compositions of active muscle AChRs are dynamic and vary producing the different subtypes of AChR and; (ii) this homeostatic plasticity allows the parasites to limit effects and habituate to the continued exposure of the anthelmintic.

4.2 Introduction

Nematode parasites are more complex than bacteria and viruses. They have a 42-700 mega-base genome (Consortium, 2019), a nervous system (Schafer, 2016) that allow learning

(Zhang et al., 2005; Ardiel & Rankin, 2010) and different motor responses to chemical stimulation (Rengarajan & Hallem, 2016). We expect that they have more complex mechanisms than viruses and bacteria to adapt and resist exposure to anthelmintic drugs.

How do nematode parasites accommodate and adapt to anthelmintic drugs designed to eliminate them? Conventional understanding of anthelmintic resistance is that it is genetically based on: i) mutation and/or selective elimination of sensitive anthelmintic targets; ii) increased drug metabolism and; iii) reduced entry and/or excretion of the anthelmintic (Beech et al., 2011;

Kotze et al., 2014). What are additional intrinsic mechanisms that allow individuals to resist effects of anthelmintics? 151

Homeostatic plasticity (Lazarević et al., 2013; Tien & Kerschensteiner, 2018) refers to the capacity of excitable cells to regulate their activity to make compensatory adjustments to long-lasting stimulation. The plasticity involves changes in ion-channels in nerves and muscles of vertebrate and invertebrate species including nematodes (Lewis et al., 1987; Turrigiano, 2012;

Bozorgmehr et al., 2013) and is triggered by changes in cytosolic calcium (Foster et al., 2018;

Padamsey et al., 2019). It has not been explored in parasitic nematodes during applications of anthelmintic drugs. The speed and level of habituation and adaptation to an anthelmintic will bear on their ability to survive and resist treatment.

We study the filarial nematode parasite, Brugia malayi (Verma et al., 2017; Kashyap et al., 2019; Verma et al., 2020). Filariae are transmitted by biting insects that feed on infected hosts’ blood and pass the parasite onto a subsequent uninfected host. A group of filarial nematodes like B. malayi that locate as adults in the host lymphatic system produces the condition known as lymphatic filariasis. They block drainage of the lymphatics, inducing gross swelling of the limbs, itching and skin infections that produce the clinical condition, elephantiasis. There are an estimated 70 million people infected with lymphatic filariasis

(Ramaiah & Ottesen, 2014) in 52 countries. There are no effective vaccines, so Mass Drug

Administration (MDA) to control and prevent infection is the only practical option. The use of

MDA gives rise to concerns about resistance and an interest in the mechanisms that the parasites use to overcome the effects of anthelmintics.

B. malayi are a tractable nematode parasite for study. They are available from an NIH- supported facility, FR3; they are amenable to knockdown; their genome is available; RNAi is a tractable procedure; their motility can be followed with Worminator and; patch-clamp recording of current responses to anthelmintic drugs can be made. In preliminary experiments we have 152 observed that exposure of B. malayi to the anthelmintic levamisole, produces paralysis which wanes over time allowing the parasite to recover motility.

Here we report quantitative motility studies, calcium fluorescence observations, patch- clamp studies and molecular observations seeking mechanistic explanations for the levamisole habituation. We followed currents in muscle cells induced by levamisole and observed that levamisole activated receptors (AChRs) over a period of 20-30 minutes: we observed a selective loss of levamisole activated currents but not acetylcholine activated currents. On the naïve muscle cell there are at least four separable types of AChRs. These are modified during maintained application of levamisole and are affected by knockdown of nra-2 that is involved in selection of protein complexes that exit the endoplasmic reticulum. We observed increases in the level of unc-38 message with a reduction in nra-2 message that drive a change in the AChR subtypes present on muscle. We conclude that the AChRs of the parasite habituate and suggest that the composition of the AChR subunits is plastic allowing the parasite to habituate to the presence the anthelmintic, levamisole. Such plasticity will support survival of nematode parasites during treatment and facilitate resistance to the anthelmintic.

4.3 Methods

4.3.1 Parasite maintenance

B. malayi adult worms were obtained from the NIH/NIAID Filariasis Research Reagent

Resource Center (FR3; College of Veterinary Medicine, University of Georgia, Athens, GA,

USA). Adult worms were maintained in non-phenol red Roswell Park Memorial Institute

(RPMI) 1640 media (Life Technologies, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS, Fisher Scientific, USA) and 1% penicillin-streptomycin (Life Technologies,

USA). The worms were stored individually in 24 well culture plates containing 2 ml of supplemented RPMI-1640 media and placed in an incubator at 37˚C supplemented with 5% CO2. 153

4.3.2 Drugs

Levamisole, acetylcholine, pyrantel, morantel, and nicotine were obtained from Sigma

Aldrich (St. Louis, MO, USA). Levamisole pyrantel, morantel, nicotine and acetylcholine were prepared in stock solutions were prepared in distilled water and diluted in recording solution to obtain final concentration. For motility assays, the recording solution was RPMI-1640, while for whole-cell recordings drugs were diluted in recording solution as mentioned below to obtain a final concentration.

4.3.3 Dissection

After dissection of the adult worms, recordings were performed at room temperature. The muscle cells and the hypodermis were exposed upon dissection by modifying the methods used for C. elegans (Richmond & Jorgensen, 1999; Qian et al., 2008). Sections of about 5mm were cut from the anterior region of the worm and placed in the recording chamber with bath solution

(23 mM NaCl, 110 mM Na acetate, 5 mM KCl, 6 mM CaCl2, 4 mM MgCl2, 5 mM HEPES,

10 mM d-glucose, and 11 mM sucrose, pH adjusted to 7.2 with NaOH, ~320 mOsmol). The base of the chamber was a 24 × 50 mm cover slip coated with a thin layer of Sylgard™. The worm section was then glued along one side using Glushield® cyanoacrylate glue (Glustitch Inc.,

Canada) thereby immobilizing it and then cut open longitudinally using a tungsten needle. The resulting ‘muscle flap’ was glued along the cut edge and the reproductive and the gut tissue were removed using fine forceps. The dissection was viewed under DIC optics (400X) using an inverted light microscope (TE2000U, Nikon, USA).

4.3.4 Whole-cell recording

Muscle flaps were incubated in 1 mg/ml collagenase (Type 1A) in bath solution for 15-

120s and washed 10 times prior to recording. The patch-clamp technique was used to record whole-cell currents from the muscle flaps as explained in (Robertson et al., 2013). Patch pipettes 154 were pulled from capillary glass (G85150T; Warner Instruments Inc., Hamden, CT, USA), fire polished and then filled with pipette solution (120 mM KCl, 20 mM KOH, 4 mM MgCl2, 5 mM

TRIS, 0.25 mM CaCl2, 4 mM NaATP, 5 mM EGTA and 36 mM sucrose (pH 7.2 with KOH),

~315-330 mOsmol). Pipettes with resistances of 3-5 MΩ were used. A 1 cm region near the tip of the electrode was covered with Sylgard™ to reduce background noise and improve frequency responses. Giga ohm seal was formed before breaking the membrane with suction. The preparation was continuously perfused in bath solution at 2 ml/min. The current signal was amplified by an Axopatch 200B amplifier (Molecular Devices, CA, USA) filtered at 2 kHz

(three-pole Bessel filter), and sampled at 25 kHz, digitized with a Digidata 1440A (Molecular

Devices, CA, USA).

4.3.5 Calcium imaging and Fluo-3 injections

Patch pipettes were filled with the pipette solution described above with added 5 µM

Fluo-3 diluted in DMSO added before each experiment and were kept in a dark environment to prevent degradation of the dye. Pipettes with resistances of 1.8-3 MΩ were used. After breaking into cells, they were left for at more than 10 minutes to allow Fluo-3 to diffuse into the muscle cell before being subjected to bath solution with added 10 mM CaCl2. Cells were consistently perfused with bath solution containing: 23 mM NaCl, 110 mM Na acetate, 5 mM KCl, 1 mM

CaCl2, 4 mM MgCl2, 5 mM HEPES, 10 mM d-glucose, and 11 mM sucrose, pH adjusted to 7.2 with NaOH, ~320 mOsmol at a rate of 1.5 mL/min.

All recordings were performed on a Nikon Eclipse TE3000 microscope (20X/0.45 Nikon

PlanFluor objective), fitted with a Photometrics Retiga R1 Camera (Photometrics, Surrey, BC,

Canada). Light control was achieved using a Lambda 10-2 two-filter wheel system with a shutter controller (Lambda Instruments, Switzerland). Filter wheel one was set on a green filter (525-

530mm) between the microscope and camera. Filter wheel two was set on the blue filter 155

(490mm) between a Lambda LS Xenon bulb light box, which delivered light via a fiber optic cable to the microscope (Lambda Instruments, Switzerland), to activate Fluo-3. Blue light emission was controlled using a shutter. Minimal exposure to blue light during recording set up was used to prevent reduction in signal strength.

Calcium signal recordings were acquired and analyzed using MetaFluor 7.10.2 (MDS

Analytical Technologies, Sunnyvale, CA, USA) with exposure settings at 250 ms with 2x binning. Maximal Ca2+ signal amplitudes (ΔF) were calculated using the equation F1-F0/F0 x

100, where F1 is the fluorescent value and F0 is the baseline value. All F0 values were determined as being the value at the time the stimulus was applied to the sample for all recordings analyzed. All pictures were taken using Ocular 2.0.1.496 (Digital optics, Auckland,

New Zealand). Exposure settings were 150 ms with 2x binning.

4.3.6 RNA extraction and cDNA synthesis

B. malayi adult worms were snap frozen and crushed into fine powder in a 1.5 mL micro- centrifuge tube using KimbleTM KontesTM Pellet PestleTM (Fisher Scientific, USA). Total RNA was extracted using TRIzol® Reagent (Life Technologies, USA) according to the manufacturer’s instructions. About 1µg of total RNA was used to synthesize cDNA using SuperScript® VILO™

Master Mix (Life Technologies, USA). Samples were either used to amplify DNA using PCR or stored at -20˚C for later use.

4.3.7 Synthesis and delivery of dsRNA

dsRNA was synthesized as explained in (McCoy et al., 2015; Verma et al., 2017). Target

PCR products were amplified using the primers in Table 1. T7 promoter sequence 5' –

TAATACGACTCACTATAG – 3' was used as an overhang to produce T7 labelled PCR products for dsRNA synthesis. Amplification was done using Techne® PRIMEG (Bibby

Scientific Limited, UK) with cycling conditions -95˚C x 5 min, 35 x (95˚C x 30s, 55˚C x 30s, 156

72˚C x 1 min), 72˚C x 10min from sequence verified cDNA templates. dsRNA was synthesized

T7 RiboMAXTM Express RNAi kit (Promega, USA) according to the manufacturer’s instructions. Concentration and purity of dsRNA were assessed using a spectrophotometer. Adult

B. malayi were soaked in RPMI media containing 30-60 µg/mL of target and control dsRNA for four days. dsRNA for lacZ was used as off-target control. Motility experiments on RNAi worms were performed after four days. Worms that were not used in the motility assay were cut into two pieces – one for electrophysiology recordings and the other was snap frozen in liquid nitrogen and stored at -80˚C for transcript analysis by qPCR.

4.3.8 Analysis of Transcript levels

cDNA from dsRNA treated worms were amplified using target and reference gene (Bma gapdh) primers (Table 1). These genes were amplified in triplicate by quantitative real-time PCR

(qPCR) using the QuantStudioTM 3 - 96 well 0.1ml Block Real time PCR detection system

(Thermofisher) and PowerUpTM SYBR® Green Supermix (Thermofisher, USA). Cycling conditions used: 95˚C x 10min, 40 x (95˚C x 10s, 55˚C x 30s). PCR efficiencies were calculated using the Design and Analysis Suite (Thermofisher, USA). Relative quantification of target gene knockdown was estimated by the ∆∆Ct method (Pfaffl, 2001).

4.3.9 B. malayi in vitro Motility Studies

Motility assays were carried out on the Worminator system in a 24-well tissue culture plate (1 worm/well) containing 1 ml of RPMI-media with L-glutamine as described (Marcelino et al., 2012). This application assess motility through pixel displacement of each worm over time, giving an output of Mean Movement Units (MMU). The more stationary the movement of the worms, the lower the motility number and the more effective the compound is on the parasite. 157

To determine the potency of levamisole on adult female B. malayi, worms were treated with various concentrations of levamisole. Worm motility was then recorded prior to the addition of levamisole and 30 seconds post treatment for each drug concentration. From this we were able generate a concentration-response curve at 30 seconds and determine IC50 values of levamisole.

Percent motility was also calculated as a percentage ratio of motility of worms after treatment at the 30 seconds time point over motility of naïve worms.

Next, to study the long-term effects of various concentrations of levamisole on adult female B. malayi motility, we conducted motility assays over a 4-hour period. Worm motility was recorded prior to the addition of levamisole, 16 seconds following the addition of levamisole and at 10, 20, 30, 40, 50, 60, 90, 120, 150, 180 and 240-minutes post treatment to generate a concentration- and time-course response analysis.

To quantify the effect of long-term application of levamisole at high concentrations, adult female B. malayi were assigned into control or drug treatment (n=4/batch) groups (1 worm/well).

Worm motility was recorded prior to the addition of levamisole, immediately following the addition of levamisole and at 30, 60, 90, 120, 150, 180, 210 and 240-minutes post treatment.

Control worms were treated with deionized water, while drug treatment worms were exposed to

100µM levamisole. Motility was recorded for 30 seconds for all time points. Three independent experiments were carried out for this study.

To investigate the effects of pyrantel, morantel and nicotine individually and in combination on adult female B. malayi motility, worms were exposed to each drug individually at a concentration of 10µM. Worms placed in the combination therapy group were exposed to

10µM of each drug (pyrantel + morantel + nicotine) simultaneously. Control worms were 158 exposed to DMSO (final concentration of 0.1% (v/v)). Worm motility was recorded as previously described above. Two independent experiments were carried out for this study.

To examine the effects of a second cholinergic agonist on worms that recovered from levamisole 4 hours post treatment, we assigned 4 worms to a control group and 20 worms to levamisole treatment. As stated previously, worm motility was recorded using the Worminator system prior to the addition the drug, immediately following addition of the drug and at 30, 60,

90, 120, 150, 180, 210 and 240-minutes post treatment. Recovered worms were maintained in the same plate containing levamisole and motility was again recorded prior to each drug treatment. Briefly, control worms were treated with DMSO, while others were treated with either

10 µM pyrantel, 10 µM morantel, 10 µM nicotine individually and in combination (pyrantel + morantel + nicotine) at 10µM each. Finally, some worms were kept as positive control

(maintained in levamisole without second exposure). Each treatment group consisted of 4 worms and the final concentration of DMSO did not exceed 0.1%. Worms were exposed to each treatment for 240 minutes and motility was recorded as stated previously.

4.3.10 Data analysis

Whole-cell patch clamp data were analyzed with Clampfit 11.1 (Molecular Devices, CA,

USA) and GraphPad Prism 5.0 software (Graphpad Software, Inc., La Jolla, CA, USA). The peak current responses from whole-cell recordings were used for analysis. For whole worm concentration–response relationships, motility/minute was plotted against log concentration.

Drug concentrations were log10 transformed before analysis. Drug concentrations were log10 transformed before analysis. The log agonist vs. response equation (variable slope) was used to generate concentration response curves to calculate EC50 values. The responses were plotted as the mean ± SE. Statistical analyses were performed on groups of values by using Two-way

ANOVA to determine whether the group means were dissimilar and Bonferroni post-hoc tests 159 were used to test significance. Paired t tests were used determine whether there were significant differences between groups where appropriate; two-tailed unpaired Student’s t-tests were used when comparing means from different preparations. GraphPad Prism 5 software was used for statistical calculations.

4.4 Results

4.4.1 Levamisole produces spastic paralysis, then a flaccid paralysis followed by recovery of motility

In the following investigations, we explore how B. malayi adapt and respond to long term applications of the cholinergic anthelmintic levamisole. Fig 4.1A shows photographs of 100 µM levamisole treated adult female B. malayi at different stages and motility plots for the B. malayi following application of different concentrations of levamisole. There were three motility phases: i) an initial spastic paralysis as the worms contracted into a ball within 1 min after application of levamisole; ii) a flaccid paralysis as the worms gradually relaxed over the next 15-

30 mins but did not move and; iii) the habituation and recovery phase as motility returned over the next 120-180 mins. These observations demonstrate that the nematode parasites adapt to exposure to an anthelmintic and that the habituation is time dependent.

Fig. 4.1B shows the concentration-dependent inhibitory effects of levamisole on worm motility during the spastic paralysis measured at 30 seconds. The IC50 of levamisole was 10 ± 2.2 nM, which highlights the potency of the effects of levamisole. Fig. 4.1C shows the concentration-independent motility response at 30 mins during the flaccid paralysis phase, which showed no significant correlation between motility and concentration of levamisole. Inspection of the worms revealed that their bodies were spread out in their respective wells but not moving at this stage. 160

We followed the motility during the recovery phase over 240 mins, where we observed that worms treated with either the highest or the lowest concentrations of levamisole recovered more rapidly, Fig 4.1D. The concentration motility response at 240 mins was U-shaped with two components: the decreasing motility with levamisole concentration explained by the spastic and flaccid paralysis associated with the opening of more AChRs with increased concentrations and; the recovery of motility that required further study. Although there was nearly full recovery when plotted in mean motility units (MMU: Worminator) with 100 µM at 4 hours, there was a detectable difference between the control and recovered pattern of the motility: the recovered worm motility was somewhat more ‘Jerky’ suggesting changes in motor control. The coefficient of variation of the motility of naïve untreated was significantly different to that of the recovered worms, Fig. 4.1E. We show later that following the recovery, the AChR channel currents are modified.

4.4.2 Calcium fluorescence and effect of levamisole

Fig. 4.2A & B shows representative traces of the effect of 30 µM levamisole and 10 mM

CaCl2 on the fluo-3 calcium signal. Note that levamisole produces an initial small rise in calcium fluorescence (Fig. 4.2A arrow: 1) that is followed by a rapid and bigger rise that starts near 250 seconds (Fig. 4.2A arrow: 2). There is then a rapid rise that reaches a peak at 22 min (1360s) before declining slowly, even in the continued presence of levamisole. Fig. 4.2B shows the response to the high 10 mM calcium bath solution which starts 15 seconds after the addition of calcium; the calcium signal rises and then is maintained until the bath is perfused with the lower calcium solution when it falls rapidly. Fig. 4.2C shows histograms of the 10% rise times, the time to peak and the amplitudes of the effects of 30 µM levamisole and 10 mM CaCl2 showing the slower rise times, times to peak and bigger amplitude of levamisole effects. The delay, the biphasic nature of the response to levamisole in contrast to the application of high calcium 161 bathing solutions suggest that the initial rise in cytosolic calcium by levamisole is mediated by the plasma membrane AChRs and the secondary larger delayed rise is mediated by a secondary intracellular release. The initial early rise in cytoplasmic calcium coincides with the phase of spastic paralysis produced by levamisole while the flaccid paralysis of levamisole occurs during the larger, delayed peak.

4.4.3 The duration of paralysis and recovery, depend on the cholinergic anthelmintic

The 4 muscle AChR subtypes, L-, P-, N- and M- AChRs of B. malayi are each preferentially activated by the cholinergic anthelmintics that they are named after: Levamisole,

Pyrantel, Nicotine and Morantel (Verma et al., 2017). We tested long duration applications of 10

µM concentrations of levamisole, pyrantel, morantel and nicotine on naïve B. malayi, Fig. 4.3A.

All agonists produced a rapid spastic paralysis that inhibited motility by 1 min that was followed by a flaccid paralysis and then a recovery of motility (habituation).

The speed of habituation was anthelmintic-dependent with nicotine showing recovery in less than 50 minutes, Fig 4.3A. The levamisole treated B. malayi recovered from the flaccid paralysis, but the recovery was slower than that of nicotine and took 80 – 240 mins. The recovery was even slower for morantel and slower again for pyrantel. Thus, the time required for habituation was anthelmintic-dependent. At 240 minutes, level of recovery was: nicotine>levamisole>morantel>pyrantel, Fig. 4.3B. The different rates of recovery of the cholinergic anthelmintics are consistent with these different anthelmintics having selective effects on the different AChR subtypes. Additionally, the lack of sustained paralysis seen over the 4-hour period for worms treated with nicotine and levamisole suggest that agonists like pyrantel and morantel, as slow-habituating anthelmintics, could have therapeutic advantages.

162

4.3.5 30 µM levamisole produces desensitization of L-AChRs with little or no effect on the other receptor subtypes

We sought to follow the habituation and adaptation to levamisole by applying a continuous concentration of 30µM levamisole under whole-cell patch-clamp conditions. Fig

4.4A shows that the inward levamisole current is not maintained and that it gradually declines over 20-100 minutes. We tested the response to a 1 min application of 100µM acetylcholine or a

1 min application of 30 µM anthelmintic before the long application of levamisole. When desensitization was complete, we again measured the responses to the application of acetylcholine or the test anthelmintic. Fig. 4.4B shows a plot of the amplitudes of the cholinergic anthelmintics currents before and after the levamisole desensitization. The levamisole currents reduced to zero (100% reduction) but the reduction in the response to acetylcholine, pyrantel, morantel and nicotine did not reach statistical significance (Fig. 4.4B, one-way ANOVA,

Bonferroni, 5 worms p > 0.05). The desensitization of the levamisole response was much greater than that of the responses to acetylcholine and the other anthelmintics.

4.4.4 100 µM levamisole inhibits/desensitizes all receptor subtypes

We then reasoned that a higher concentration of levamisole, 100µM applied for a longer duration, would have a less selective inhibitory effect on the nAChRs than 30 µM levamisole.

We performed electrophysiology on the body wall muscle cells under whole-cell patch-clamp in

5 different recovered B. malayi female worms after 4 hours of long-term 100 µM levamisole exposure. The recordings made in the presence of 100µM levamisole showed significantly reduced currents for all the AChR agonists tested, Fig. 4.5. The percent reduction potency series was levamisole (100%) > ACh (78%) > morantel (74%) > pyrantel (66%) > nicotine (48%). The high concentration of levamisole blocked all L-AChR responses to levamisole but responses to 163 the other agonists of the other AChR subtypes remained, although reduced. These remaining

AChR subtypes will be available to contribute to the recovery of motility.

4.4.5 unc-38 is upregulated, and nra-2 is downregulated, in levamisole habituated Brugia

We have seen that adult female B. malayi treated with 100µM levamisole adapts, allowing motility to recover after 4 hours, although with a slightly modified motility phenotype.

We looked for changes in expression levels of the AChR subunits: unc-38, unc-29, unc-63, acr-

8, acr-16 and acr-26. We hypothesized that expression of one or more of these subunits would change and be involved in the habituation. We observed that in the recovered worm, the unc-38 transcript was upregulated, while the expression of the other genes was unaffected. We also observed that nra-2 (a gene that encodes for an ER retention protein) was downregulated, Fig.

4.6A. nra-2 in C. elegans that leads to faster adaptation to levamisole (Almedom et al., 2009;

Kamat et al., 2014). These observations indicate a role for nra-2 and unc-38 in part of the habituation response to levamisole.

4.4.6 nra-2 knockdown speeds but unc-38 knockdown slows levamisole desensitization/recovery

To determine the effect of nra-2 on the adaptive response to levamisole, we knocked down the nra-2 transcript by soaking the worms with dsRNA for four days. We then found that these worms recovered faster than control lacZ worms in 100µM levamisole, Fig. 4.6B. qPCR on these worms confirmed knockdown of the nra-2 transcript, Fig. 4.6C. We also knocked down unc-38 transcript and found that these worms recovered more slowly from levamisole and recovery took longer, Fig. 4.6B. The RNAi experiments and expression changes of unc-38 and nra-2 indicate that these two genes are key components of the habituation response of B. malayi to levamisole.

164

4.4.7 nra-2 knockdown reduces levamisole currents but not ACh currents

To examine the effects of nra-2 further, we made whole cell patch-clamp recordings from muscle cells treated with dsRNA for 4 days to knockdown nra-2, Fig. 4.7A. The muscle cells were each given a standard perfusion for 25s with 30 µM concentrations of acetylcholine, levamisole, pyrantel, morantel, and additionally bephenium to look for any appearance of B-

AChRs (Qian et al., 2006). We measured the mean peak currents measured in 5 separate preparations in LacZ treated control worms and dsRNA nra-2 treated worms. There was a significant reduction in the amplitude of the levamisole currents that was statistically significant but not with the other cholinergic anthelmintics, Fig. 4.7B. Again, we see evidence of agonist selective effects of nra-2 knockdown on the response to levamisole. The reduction in the levamisole sensitivity on the muscle show that nra-2 is required for maintaining and fostering the

L-AChRs.

4.4.8 Reversal of levamisole habituation and nra-2 expression levels after removing levamisole

We looked to see if there was a reversal of habituation after removal of levamisole. We treated B. malayi with 100 µM levamisole for 4 hours as before. We then transferred these worms to fresh media without levamisole and then tested groups of 4 worms with 100 µM levamisole after an interval of 1, 30, 60, 90 or 120 minutes, Fig. 4.8A&B. The worms re-tested with levamisole 1 min after habituation in the fresh media (t1: 1 min) showed little response to levamisole with only a 27.8 ±15.8%, reduction in motility, Fig. 4.8B&C. The worms tested with levamisole after 120 minutes (t120) in fresh media responded well to the second exposure to levamisole with a 70.65 ± 7.59% reduction in motility, Fig. 4.8B&C. Fig. 4.8C shows that the inhibition of motility with the t-1 worms was significantly smaller than the t-120 worms. 165

We also investigated the change of expression of nra-2 and unc-38 following return to fresh levamisole-free media, to determine if there were expression changes associated with the loss of this habituation. The transcript levels were measured in: i) naïve untreated worms; ii) in control worms habituated after four hours in levamisole and; iii) in worms habituated for four hours and then moved to fresh RPMI media for 120 minutes. The expression level of nra-2 was decreased and unc-38, was increased in habituated worms, Fig. 4.8D, as before (see Fig. 4.6A).

However, the nra-2 transcript level of worms removed from levamisole and soaked in fresh media for 2 hours (4hr+2hr) returned to the baseline level, but the unc-38 expression appeared to be unchanged at this time, Fig 4.8D. Here, we see again the correlation between the response to levamisole and expression of nra-2.

4.4.9 Recovery and Knockdown of AChR receptor genes

Fig.4.9A shows that the knockdown of single nAChR subunit genes (unc-38, unc-63, acr-

8, acr-26 and acr-16) had little detectable effect of the spontaneous motility of B. malayi.

Knockdown of unc-29 significantly reduced but did not abolish spontaneous motility of worms.

This was expected as UNC-29 and UNC-38 together were shown to be necessary for spontaneous motility (Verma et al., 2017): UNC-29 is the only non-alpha subunit in body muscle, so the effect of knocking it down was anticipated to have a bigger impact on motility than knocking down unc-38, one of five alpha subunits. Regardless, all these worms showed sensitivity to levamisole and were paralyzed upon treatment with 100 µM levamisole (Fig.

4.9A).

When we looked at recovery however, we found that knockdown of unc-38, acr-26, and acr-16 delayed recovery, and unc-63, unc-29 and acr-8 did not, Fig 4.9A. The knockdown of unc-38, acr-26 and acr-16 together had a bigger effect and prevented (>12 hours) rather than just delayed recovery, Fig. 4.9B. Thus, the genes that are essential for normal levamisole recovery 166

(unc-38, acr-26, and acr-16) are not essential for normal motility (unc-29). There is a change from the genes that are essential for normal spontaneous motility to the genes that are essential for the recovery of motility (habituation) to levamisole. This change is associated with the modified motility phenotype, Fig 4.1E.

4.5 Discussion

4.5.1 Dynamic and Plastic AChRs in Brugia muscle

Here we have documented more complex time-dependent and concentration-dependent responses of a parasitic nematode to an anthelmintic. These observations illustrate some of the limitations of simple anthelmintic concentration-response plots when evaluating efficacy of these drugs. The anthelmintic we studied was levamisole, which is a selective agonist of specific subtypes of nematode nicotinic AChRs in Brugia malayi. In the C. elegans model nematode levamisole also produces time-dependent responses (Lewis et al., 1987) but there are only two nicotinic AChRs present on muscle: the nicotine-sensitive AChR and the levamisole-sensitive

AChR (Richmond & Jorgensen, 1999).

In contrast, there are at least four receptor subtypes types on Brugia malayi muscle, Fig.

6.9 referred to as the L-, P-, M- and N- subtypes (Verma et al., 2017). Levamisole preferentially activates L-AChRs. Parasite muscle AChR subtypes are produced by different pentameric combinations of nAChR subunits that include UNC-38, UNC-29, UNC-63, ACR-8, ACR-16 and

ACR-26. The AChR subunits combine in different pentameric and stoichiometric arrangements to produce the AChR subtypes that have distinctive pharmacological properties and sensitivities to levamisole (Boulin, 2011; Boulin et al., 2011; Buxton et al., 2014; Williams et al., 2020).

The numbers of the different AChR subtypes will alter whole muscle current responses and muscle responses to different cholinergic anthelmintics. B. malayi muscle responses are characterized by an initial spastic paralysis, followed by flaccid paralysis and then a gradual 167 recovery of motility over 4 hours. We see desensitization (decline) of the levamisole current response when 30 µM levamisole is applied for a long time (~20 minutes) while current responses to acetylcholine remain are not desensitized. Long term (4 hours) application of levamisole increases unc-38 and decreases nra-2 expression; nra-2 knockdown also selectively reduces levamisole current responses, Fig. 4.7B. In C. elegans AChRs sensitive to levamisole are also decreased and AChRs insensitive to levamisole are increased in nra-2 null mutants

(Almedom et al., 2009). These observations reveal dynamic mechanisms of AChRs in parasitic nematodes by which responses to levamisole are modified allowing adaptation to limit effects of levamisole.

What is the functional significance of the plastic and different nAChR subtypes on muscle cells? The nAChR subtypes are all non-selective acetylcholine-gated ion-channels that differ in their calcium permeability, sensitivity to acetylcholine, rate of desensitization and perhaps membrane location. The levamisole-preferring, L-AChR, is made up of the subunits of

UNC-38, UNC-63, ACR-8, and UNC-29 and is 20x more permeable to calcium than channels composed of UNC-63, ACR-8 and UNC-29 subunits (Buxton et al., 2014). The AChR channel that desensitizes faster than others, is the N-AChR, that is composed of ACR-16 subunits

(Abongwa et al., 2016) and that is selectively activated by nicotine and acetylcholine rather than levamisole, pyrantel and morantel. The effects of nicotine on motility desensitizes faster than levamisole which in turn desensitizes faster than morantel and pyrantel, Fig. 4.4A.

Fig. 4.10 illustrates a summary diagram with the different subtypes of nAChRs found on

B. malayi muscle membrane: it shows the editing of the AChR subtypes by NRA-2 inhibiting the insertion of levamisole insensitive AChRs into the plasma membrane. Fig. 4.10 also illustrates how the increased cytosolic calcium that we have observed with prolonged levamisole 168 application may promote unc-38 expression, inhibit nra-2 expression, inhibit motility, interact with TRP channels (Ramsey et al., 2006; Kashyap et al. 2019) and contribute to AChR desensitization.

The general processes of homeostatic plasticity are well recognized in neuroscience and refer to the capacity of excitable cells to regulate their own activity. Intracellular calcium is the primary coordinator that regulates their excitability with compensatory adjustments occurring over different time scales. The plasticity involves changes in the number and distribution of membrane ligand-gated ion-channels, voltage-activated ion-channels, calmodulin, calcineurin, intracellular calcium release and uptake (Chen et al., 2008; Ge et al., 2009; Turrigiano, 2012). In

C. elegans the plasticity is associated with regulation of the number of AChRs presented to the plasma membrane (Jensen et al., 2012) and with AChR subunit composition and transfer to the plasma membrane being regulated by NRA-2 (Almedom et al., 2009). The homeostatic plasticity will allow the parasite to control and adapt motility to different environments that the nematode finds itself in as it moves through its life cycle and around in the host. In some environments, it will need to more rapid contractions for swimming, in others it will need a slower sustained contraction to hold onto its location. It will also allow the parasite to adapt to the cholinergic anthelmintics as we have seen.

4.5.2 Desensitization and flaccid paralysis

The effects of levamisole on motility are time- and concentration-dependent and characterized by the initial spastic paralysis phase (<20min) due to the opening of nAChR channels on body muscle; this is followed by a phase of flaccid paralysis as the body muscles relax (from the end of spastic paralysis to <4hrs), despite elevated cytosolic calcium levels, and

AChR channel desensitization. The features of mammalian nicotinic receptor channel desensitization have been studied extensively (Huganir & Greengard, 1990; Quick & Lester, 169

2002; Dani, 2015) and involve different mechanisms including slow adjustments of the channel amino-acid positions and changes is receptor phosphorylation driven by calcium dependent kinases and phosphatases. The regulators of desensitization of AChRs that present in C. elegans muscle and Brugia include: TAX-6, a calcineurin A subunit, that affect desensitization of AChRs in rat chromaffin cells (Khiroug et al., 1998); SOC-1 (a multi-subunit adaptor protein); and PLK-

2 (a serine/threonine kinase (Khiroug et al., 1998; Gottschalk et al., 2005). We have not addressed these regulators in this study, but they are anticipated to affect the rate of levamisole desensitization and homeostatic plasticity.

4.5.3 Calcium homeostasis

Maintained application of levamisole, Fig. 4.2, produces an initial modest increase in cytosolic calcium followed by secondary larger increase in cytosolic calcium, Fig. 4.10.

Ryanodine receptors (RyRs: UNC-68) are present in nematode parasite muscles

(Ascaris:(Puttachary et al., 2010; Robertson et al., 2010); Brugia: Wormbase) as well as C. elegans, UNC-68: Wormbase; (Maryon et al., 1996; Hamada et al., 2002). The rise in cytosolic calcium following opening of AChRs by levamisole then produces muscle contraction mediated by the calcium contraction coupling pathway (Robertson et al., 2010; Gao & Zhen, 2011; Hwang et al., 2016). TRP channels (TRP-2, GON-2 and CED-11) channels present in Brugia muscle also contribute to the muscle contraction (Feng et al., 2006; Trailovic et al., 2008; Verma et al.,

2020). TRP channels are subject to regulation by intracellular messengers including direct or indirect activation and inhibition by calcium and PKC, (Yao et al., 2005; Feng et al., 2006;

Hasan & Zhang, 2018), Fig 4.10. Inhibition of the TRPs will contribute to calcium homeostasis as will calcium uptake: i) ER and plasma membrane calcium ATPase (SERCA: sca-1a & sca-1b:

(Cho et al., 2000); ii) Na/Ca exchangers; iii) Na/Ca/K exchangers; and iv) Ca/cation exchangers, ncx-1 to ncx-10 (Sharma et al., 2013). 170

Our calcium fluorescence experiments, Fig. 4.2, indicate that cytosolic calcium remains elevated during the earlier part of the flaccid paralysis indicating that that calcium homeostasis is not the explanation for the flaccid paralysis. One explanation that allows a relaxation despite elevated calcium is a calcium-induced calcium-desensitization of the myosin light chain by dephosphorylation with myosin light chain phosphatase (Rembold, 1992; Murahashi et al., 1999;

Khromov et al., 2006). Fig. 4.5 shows that application of 100µM levamisole for 4 hours blocks the levamisole currents and inhibits but does not block currents from of other AChRs. These remaining AChRs, once the high calcium concentration of sarcoplasm has been cleared by the homeostatic mechanisms (Clapham, 2007; Martin & Richmond, 2018) could permit the return of motility as the sensitivity of myosin light chain kinase returns as the excess cytoplasmic calcium is removed. Inhibition of the homeostatic mechanisms is anticipated to inhibit the return of motility.

4.5.4 Gene regulation and Recovery

We tested for the recovery from levamisole desensitization following removal of the levamisole, Fig. 4.8. We find that there was a gradual return of sensitivity over 4 hours associated with a return towards control nra-2 message levels but with unc-38 levels remaining elevated. NRA-2 is expressed in endoplasmic reticulum of body wall muscle in C. elegans where it controls the composition of different AChR subunits in the plasma membrane so that nra-2 null mutants are less sensitive to levamisole. NRA-2 is a homolog of human nicalin that contains a predicted calcium-sensitive EF hand that extends into the lumen of the endoplasmic reticulum suggesting actions related to emptying of calcium from the ER.

4.5.5 Therapeutic significance

We have seen that B. malayi can adapt to the anthelmintic levamisole by desensitization of the levamisole sensitive receptors (L-AChRs) on muscles. During treatment of an infected 171 host with levamisole there will be gradual rather than a rapid rise in the concentration of levamisole at the site of the worm. This time-dependent increase in levamisole concentration at the site of the worm can permit adaptation and homeostatic plasticity processes that we have described to allow survival and resistance to treatment.

The desensitization and homeostatic plasticity mechanisms that we have observed are driven by changes by increases in intracellular calcium and could also be at the heart of the transient effect of diethylcarbamazine (Verma et al., 2020). For these drugs to be effective particularly against nematode parasites that are less sensitive to drugs like levamisole and diethylcarbamazine, then repeated dosing allowing some recovery from the desensitization may enhance a therapeutic benefit (Mak & Zaman, 1980).

4.6 Conclusion

We have seen that the response to an anthelmintic levamisole is time dependent as the parasite adapts to the presence of the drug. There is a spastic paralysis, a flaccid paralysis, and then recovery and change in nAChR subtypes present on the muscle cells. It is proposed that the nAChR subtypes are dynamic, showing a homeostatic plasticity and that activity and cholinergic anthelmintics drive the subunit composition and thereby change the nAChR subtype. Such a mechanism will allow the nematode parasite to accommodate to an environmental change as well as exposure to an anthelmintic and thereby resist treatment.

4.7 Acknowledgements

We would like to thank NIH/NIAID Filariasis Research Reagent Repository Center

(www.filariasiscenter.org) for regular supply of adult and microfilariae of Brugia malayi worms.

The research was funded by The NIH NIAID Grants R01AI047194 and R21AI138967 to R.J.M. and The EA Benbrook Endowed Chair of Pathology and Parasitology. The funding agencies had no role in the design, execution or publication of this study. 172

The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infectious Diseases

4.8 Author contributions

S.S.K., S.V., M.M., M.W., P.W., A.P.R., and R.J.M. conceived the research idea and hypothesis; S.S.K. performed dsRNA, qPCR, some of the motility experiments; S.V. conducted patch clamp experiments; M.M performed some of the motility experiments (Figures 4.1, 4.3 and

4.8); M.W. conducted some of the motility experiments at the preliminary stages of this study;

P.W. performed calcium imaging experiments; S.S.K., S.V., M.M., P.W., A.P.R., and R.J.M wrote, revised and edited the manuscript.

4.9 Competing interests

The authors declare no competing interests.

4.10 Figure and Table Legends

Table 4.1. List of primers used in this study

Primer Name Description Sequence 5' - 3'

nra-2f Bma nra-2 dsRNA 5’ GCAGCAATTGTTAAATGCAA

nra-2r Bma nra-2 dsRNA 3’ ATCCTGGTATTGCTGAATGG

unc-38f Bma unc-38 dsRNA 5’ TACTATCCGTCCGTCGAGTG

unc-38r Bma unc-38 dsRNA 3’ TTCACCACTATGCGATGGTA

gapdhf Bma gapdh dsRNA 5' GACGCTTCAAGGGAAGTGTTTCTG

gapdhr Bma gapdh dsRNA 3' GTTTTGGCCAGCACCACGAC

LacZf LacZ dsRNA 5' CGTAATCATGGTCATAGCTGTTTC

LacZr LacZ dsRNA 3' CTTTTGCTGGCCTTTTGCTC

173

Table 4.1 Continued

Primer Name Description Sequence 5' - 3'

acr-16f Bma acr-16 dsRNA 5' CGACCAGGAGTTCATCTCTC

acr-16r Bma acr-16 dsRNA 3' GAAATTGGGCTCTTTCCATT

acr-26f Bma acr-26 dsRNA 5' CTCAATTAAATTCGGCTCGT

acr-26r Bma acr-26 dsRNA 5' AGCGTCTTCCGTCTGATATG

unc-63f Bma unc-63 dsRNA 5' CAGAAACATTGCTTGGCTTT

unc-63r Bma unc-63 dsRNA 3' AGGTGATTCACAGCATGGAT

unc-29f Bma unc-29 dsRNA 5' CCTCATCCACAATCCCACTA

unc-29r Bma unc-29 dsRNA 3' CGGTTTTGGTCTTTGCATAC

acr-8f Bma acr-8 dsRNA 5' CGGTTTCCAAATTGATGTTC

acr-8r Bma acr-8 dsRNA 3' AGGATACAGGCGTTCATGTC

174

Figure 4.1. Levamisole is highly potent in paralyzing adult B. malayi females, but the worms undergo desensitization within four hours. A) Concentration- and time-dependent analysis of female B. malayi motility following four hours exposure to various concentrations of levamisole; n=12 worms per concentration over three biological replicate studies. B) Concentration-response analysis of adult B. malayi females treated with various concentrations of levamisole. Motility was measured 30sec post levamisole treatment. IC50 = 10 ± 2.2 nM; n=8 per concentration over two biological replicate studies. Representative image showing the spastic paralysis phenotype. 175

C) Concentration-response analysis of B. malayi adult females treated with various concentrations of levamisole. Mean motility unit (MMU) values were taken at the 30 mins time point post levamisole treatment; n=12 worms per concentration over three biological replicate studies. Representative image showing the flaccid paralysis phenotype. D) Concentration- response analysis of B. malayi adult females treated with various concentrations of levamisole. MMU values were taken at the 240 mins time point post levamisole treatment; n=12 worms per concentration over three biological replicate studies. Representative image showing the recovery of the motility phenotype. E) Percent co-efficient of variation in naïve and recovered worms at t240; n=60 over 15 biological replicate studies. Student’s t-test was performed for statistical analysis and p<0.05 was considered as statistically significant.

176

Figure 4.2. Long-term Levamisole application generates a calcium response in Brugia malayi muscles injected with Fluo-3. A) Representative trace of a calcium signal to long-term application of 30 µM Levamisole. Arrow 1 represents the first initial increase in the calcium signal. Arrow 2 indicates the second, much larger increase which plateaus and remains until removal of the stimulus. Yellow box indicates levamisole application. B) Representative response to 10 mM CaCl2. Arrow represents the initiation of the response. Grey box represents stimulus application. C) Quantification of the average time taken for the calcium signal to reach 177

10% of the overall amplitude in response to 10 mM CaCl2 (black bar) and 30 µM levamisole (White bar). * Significantly different to CaCl2 (CaCl2 vs lev P = < 0.0181, t = 2.961, df = 8, unpaired t-test). D) Quantification of the average time for the calcium signal to reach peak (100%) in response to 10 mM CaCl2 (Black bar) and 30 µM levamisole (White bar). * Significantly different to CaCl2 (CaCl2 vs Lev P = < 0.0134, t = 3.160, df = 8, unpaired t-test. E) 2+ Amplitudes of Ca signals in the muscle in response to 10 mM CaCl2 (Black bar) and 30 µM levamisole (White bar). N.S. not significantly different to CaCl2 (CaCl2 vs Lev P = 0.4622, t = 0.7723, df = 8, unpaired t-test). All values represented as mean ±SEM. CaCl2 recordings n = 5 muscles from 5 individual worms. Levamisole recordings n = 5 muscles from 5 individual worms.

Figure 4.3. Adult female B. malayi possess multiple nAChR subtypes that vary in sensitivity to various nAChR agonists. A) Concentration- and time-dependent analysis of the inhibitory effects of nicotine (brown), levamisole (blue), morantel (red), and pyrantel (green) on worm motility. Worms were treated at a concentration of 10µM for each drug individually. N = 7 worms per concentration over two biological replicate studies. B) Bar chart (mean ± SEM) showing the mean motility of B. malayi after 4hrs (240 mins) post agonists treatment. The efficacy in inhibition of motility at 240 mins was as follows: 10µM pyrantel > 10µM morantel >10µM levamisole > 10µM nicotine; n = 7 worms per concentration over two biological replicate studies. One-way ANOVA was performed for statistical analysis and P<0.05 (*), P < 0.001 (***) and P< 0.0001 (****) were considered statistically significant. 178

Figure 4.4. B. malayi muscle cells desensitize to continuous levamisole exposure under whole- cell patch-clamp without losing sensitivity to other nicotinic agonists. A) Representative whole- cell patch-clamp recordings demonstrating cells desensitize with loss of inward current, 15-30 mins after continuous application of 30µM levamisole. Inward currents were observed when the same preparation was exposed to nAChR agonists (30 µM ACh, pyrantel, nicotine and morantel, respectively). B) Bar chart demonstrating inward current before and after levamisole 179 desensitization. Note only pyrantel and morantel showed a significant reduction, paired t-test, n=5 from 5 different worms, p < 0.05.

Figure 4.5. B. malayi whole worms after long term whole worm levamisole exposure loses agonist sensitivity in muscles under patch-clamp. A) Representative single electrode patch-clamp recording from muscle cells of control B. malayi with no prior levamisole exposure demonstrating inward current produced by different nicotinic agonists (ACh, pyrantel, nicotine, and morantel) each at 30 µM concentration for 30 seconds. B) Representative single electrode patch-clamp recording from B. malayi muscle cells after 4hour 100 µM levamisole exposure shows reduction inward currents to different nicotinic agonists (ACh, pyrantel, nicotine and morantel) each at 30 µM concentration for 30 seconds in the presence of 100 µM levamisole. C) Bar chart demonstrating comparative inward currents in muscle cells treated with 30 µM ACh, pyrantel, nicotine and morantel in: control vs the worms maintained in 100 µM levamisole for 4 hours (paired t-test, n=6, p<0.001; p<0.05 for nicotine). 180

Figure 4.6. unc-38 and nra-2 transcript levels altered during levamisole recovery and nra-2 knockdown results in faster recovery in adult female B. malayi. A) Transcript level analysis on recovered worms after being incubated in levamisole for four hours. unc-38 was significantly upregulated while nra-2 was significantly downregulated (Student’s t-test; *** - p<0.0005, * - p<0.05) whereas the other transcripts tested, unc-29, unc-63, acr-8, acr-16 and acr-26 were unaltered. N=12. B) Knockdown of nra-2 using dsRNA significantly increases the rate of recovery (Two-way ANOVA; p<0.05), whereas knockdown of unc-38 transcript resulted in the worms not completely recovering after four hours (Students t-test; p<0.005). lacZ dsRNA was used as non- specific control and control worms were soaked in water. C) Knockdown of unc-38 and nra-2 transcripts after four days of incubation in dsRNA assayed using qPCR. Knockdown in the unc- 181

38 transcript was 77.27 ± 7.21% and nra-2 was 91.97 ± 2.57%). The knockdown of these transcripts in worms soaked in non-specific lacZ control was – unc-38 – 15.87 ± 2.69% and nra-2 – 14.23 ± 2.32%. N = 5 worms over two biological replicate studies.

Figure 4.7. Whole-cell patch-clamp recording from muscle cells of the nra-2 dsRNA treated adult worms. A) Representative whole-cell patch-clamp recordings demonstrating that adult worms soaked in LacZ dsRNA treated worms produced inward currents (30 µM ACh, pyrantel, bephenium and morantel, respectively) and the inward current response to levamisole and pyrantel were significantly reduced in nra-2 dsRNA treated worms. B) Histogram demonstrating inward current response to (30 µM ACh, pyrantel, bephenium, and morantel) in LacZ and nra-2 dsRNA treated worms. The responses to levamisole and pyrantel were significantly reduced (paired t-test, p<0.005, n= 5 cells from 5 worms).

182

Figure 4.8. Treated adult female B. malayi become re-sensitive to levamisole after 120 mins in RPMI media. A) Shows images of worms under spastic paralysis for t-0, t-1 and t-120 worms. B) Worms treated in levamisole recover after four hours (t-0; grey). These worms were transferred into RPMI media and treated with a second dose of levamisole after 1 minute (t-1; blue) and 120 minutes (t-120; purple). Control worms (black) were treated with water. C) Histogram of the spastic paralysis effect of levamisole on the above worms. Inhibition of motility in t-1 worms were significantly lower compared to the t-120 worms (Student’s t-test; p<0.005; n=8 over two biological replicates). D) Transcript levels of nra-2 in worms recovered under levamisole after the first four hours (t-0 worms) and in worms moved to fresh RPMI media for 120 minutes (t-120). N=6 over two biological replicates. 183

Figure 4.9. UNC-38, ACR-26 and ACR-16 are required for recovery of motility with 100 µM levamisole treatment (habituation). A) Shows the effect of knockdown on unc-38, unc-63, unc-29, acr-8, acr-16 and acr-26 on levamisole habituation. Knockdown of unc-38, acr-16 and acr-26 caused significant impairment in levamisole habituation at t-240 minutes. B) Triple knockdown of unc-38, acr-26 and acr-16 resulted in no reduction in motility in naïve worms, but the levamisole habituation phenotype was severely impaired (2-way ANOVA, p<0.05, n=8 over two biological replicates).

184

Figure 4.10. Summary diagram of proposed mechanisms of homeostatic plasticity allowing levamisole habituation. Levamisole preferentially opens the L-subtype of AChRs, depolarization and entry of calcium. The calcium influx in the muscle produces contraction and spastic paralysis. The L-subtype of AChRs desensitizes over ~20 minutes (1: Desensitization). Calcium also builds up in the cell with the support of the calcium-induced calcium release via the ryanodine receptor (RyR). The sarcoplasmic calcium then gradually decreases due to homeostatic uptake mechanisms (2: Ca Homeostasis) that include the sarcoplasmic endoplasmic reticulum calcium ATPase (SERCA). unc-38 is upregulated while nra-2 is down regulated (3: Gene Regulation) that leads to a dis-inhibition of insensitive levamisole AChRs (iL-) in the ER containing UNC-38, ACR-26 and ACR-16 (4: Receptor translocation). The iLs are then translocated to the membrane facilitation the recovery of motility of the worms. 4.11 References

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CHAPTER 5. GENERAL CONCLUSION

Anthelmintic resistance is a global phenomenon, with parasites inflicting severe morbidity and mortality on humans and animals. For several years, researchers have taken the approach of trying to elucidate the mechanisms of resistance and gain meaningful understanding of parasitic diseases. Although some level of progress has been made, the problem of anthelmintic resistance still lingers due to gaps in our understanding of parasite biology, inability to culture and reproduce the life cycle stages of the parasites in vitro, lack of completely sequenced and accurately annotated genomes, along with other challenges. Despite these setbacks my PhD studies provide strategies that can be employed in the fight against resistance.

We chose the nematode intestine as our site of investigation due to its importance to the parasite, and the fact that numerous studies have focused mainly on muscle and neuronal cells as target sites for anthelmintics. The nematode intestine is involved in the secretion of enzymes, absorption of digested nutrients, provides an efficient barrier to the luminal environment, and synthesizes molecules that may potentially influence the host immune response. Therefore, impairment of this essential organ can be detrimental to the parasite’s survival, thus making it a suitable target tissue for anthelmintic therapy. Our studies have shown subcellular localization of four nAChR subunit mRNAs that constitute the putative levamisole receptor in the intestine of A. suum. We were also able to successfully demonstrate that the intestine is responsive to a cholinergic anthelmintic, levamisole, which validates the presence of functional nAChR receptors in A. suum intestine. The role of these receptors is still left to be explored and may involve some paracrine function. However, these findings highlight the diverse distribution of nAChR in parasitic nematodes, which are not restricted to excitable tissues, but also non-

192 excitable tissues. Therefore, this creates new avenues for future studies that do not only focus on nematode muscle and neurons, but also the intestinal epithelium as a target for new therapies.

In our next study, we were able to demonstrate that the avermectins, namely abamectin and ivermectin and the milbemycin, moxidectin exhibited positive allosteric modulator (PAMs) effects on the O. dentatum levamisole nAChR. In contrast, both avermectins acted as negative allosteric modulators (NAMs) on the pyrantel/tribdendimidine subtype, while moxidectin maintained its PAM effect. These results show that the macrocyclic lactones mode of action is not restricted to GluCls but also nAChRs. Hence, it provides knowledge on at least one mechanism in which combination therapy may enhance drug efficacy and delay the onset of anthelmintic resistance.

Finally, we were able to provide a mechanism by which Brugia malayi worms were able to adapt to levamisole exposure by regaining motility after initial paralysis. Through various experiments, these studies reveal that the nematode muscle nAChRs are dynamic and have the capacity to produce receptor subtypes that are insensitive to levamisole within a span of less than

4 hours at the muscle surface, thus aiding in recovery. These findings gives us insight of a mechanism of resistance used by parasitic nematodes which might be counteracted by use of combination therapy leading to the parasite’s demise.

5.1 Future directions

Since the nematode intestine has been proposed to be a suitable target for anthelmintic therapy, investigation into its sensitivity to other anthelmintics, not limited to cholinergic agonists would be desirable. This is because there is existing RNAseq data that suggest the presence of various types of ion channels such as GluCls, GABA and other channels which may be targets for a wide range of currently existing and novel anthelmintics. Additionally, Cry5B 193 which is widely known to act on the nematode intestine, could also be investigated more carefully in vitro along with the employment of combination therapy to improve drug efficacy.

The PAM and NAM effects exhibited by the macrocyclic lactones could be further explored on parasites in vitro, specifically Oesophagostomum dentatum, Brugia malayi and other parasitic nematodes whose motility can be analyzed and quantified. Hence, using the knowledge gained from the pharmacology studies with the levamisole receptors being allosterically modulated by macrocyclic lactones, it is likely that combination therapy between cholinergic agonists and the macrocyclic lactones could abolish receptor desensitization and facilitate sustained paralysis.