Gap Junctions in the Mosquito, Aedes Aegypti DISSERTATION Presented

Gap Junctions in the Mosquito, Aedes aegypti

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Travis Calkins

Graduate Program in Entomology

The Ohio State University

2017

Dissertation Committee:

Dr. Peter Piermarini, Advisor

Dr. David Denlinger

Dr. Carol Anelli

Dr. Reed Johnson

Copyrighted by

Travis Lee Calkins

2017

Abstract

Mosquitoes are the most dangerous animals on the planet due to the pathogens they transmit to humans. The yellow fever mosquito, Aedes aegypti, is the study organism of this dissertation, as well as the primary vector for the viruses that cause yellow, dengue, chikungunya, and Zika fevers in humans. Unfortunately, many of these diseases lack effective vaccinations and/or therapeutics and instead must be prevented through control of the mosquito vector. Our current approach to mosquito control relies primarily on the use of insecticides to suppress mosquito populations. While these chemicals are incredibly effective at killing mosquitoes, they also exert a strong selective pressure driving the evolution of resistance. In order to combat this resistance, current compounds must be modified and new targets need to be identified. In this dissertation

I focus on the latter with mosquito gap junctions as my potential targets of interest. Gap junctions are intercellular channels that mediate direct communication between adjacent cells via the transfer of small molecules and/or ions. Gap junctions are formed by protein subunits known as connexins in vertebrates and innexins in invertebrates, which are evolutionarily distinct sharing no significant amino acid homology. Despite their distinct evolutionary origins, both connexin and innexin formed gap junctions play integral roles in processes from embryogenesis to reproduction. Here I expand the knowledge base of mosquito gap junctions through examination of gene expression, ii protein localization, pharmacological inhibition, and RNAi based gene knockdown. I find that gap junction inhibitors kill and incapacitate larval and adult female mosquitoes, and inhibit diuresis in adult females. Moreover, RNAi to knockdown the mRNA expression of innexin genes decreases survival. Additionally, I show splice variation for 2 innexin mRNAs, describe innexin gene expression throughout the animal, localize the inx3 protein, and characterize changes in innexin gene expression after a blood meal. Finally,

I demonstrate the involvement of innexins in crop muscle contractions and the potential cell signaling mechanism that leads to the opening or closure of gap junctions in the crop musculature. Taken together this work sets a strong foundation for future investigations of gap junctions in the mosquito and suggests that gap junctions may provide targets for the development of novel insecticides.

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Acknowledgments

I thank my advisor Dr. Peter Piermarini for everything he has contributed to this project, my education, and my career, without his guidance none of this would have been possible. I also thank my committee members, Drs. David Denlinger, Carol Anelli, and

Reed Johnson, for their critical analysis and helpful guidance throughout my program.

Finally I thank all of the people who have supported me and fueled my love of science along the way.

Vita

2012 ...... B.A. Biology, The College of Wooster

2013 to Present ...... Graduate Research/Teaching Associate,

Department of Entomology, The Ohio State

University

Publications

Yang, Z., B.M. Statler, T.L. Calkins, E. Alfaro, C.J. Esquivel, M.F. Rouhier, J.S. Denton, and P.M. Piermarini. 2016. Dynamic expression of genes encoding subunits of inward rectifier potassium (Kir) channels in the yellow fever mosquito Aedes aegypti. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology. 204, 35-44.

Lehtinen, R.M., T.L. Calkins, A.M. Novick, J.L. McQuigg. 2016. Re-assessing the conservation status of an island endemic frog. Journal of Herpetology, 50(2): 249-255. doi: http://dx.doi.org/10.1670/14-161

Calkins, T.L. and P.M. Piermarini. 2015. Pharmacological and genetic evidence for gap junctions as potential new insecticide targets in the yellow fever mosquito, Aedes aegypti. PLoS One. doi: 10.1371/journal.pone.013708

Piermarini, P.M., S.M. Dunemann, M.F. Rouhier, T.L. Calkins, R. Raphemot, J.S. Denton, R.M. Hine, and K.W. Beyenbach. 2015. Localization and role of inward rectifier K+

v channels in Malpighian tubules of the yellow fever mosquito Aedes aegypti. Insect Biochem. Mol. Biol. 67: 59-73, 2.

Calkins, T.L., Woods-Acevedo, M.A., Hildebrandt, O., & Piermarini, P.M. 2015. The molecular and immunochemical expression of innexins in the yellow fever mosquito, Aedes aegypti: Insights into putative life stage- and tissue-specific functions of gap junctions. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 183, 11–21. doi:10.1016/j.cbpb.2014.11.013

Piermarini, P.M., and T.L. Calkins. 2014. Evidence for intercellular communication in mosquito renal tubules: A putative role of gap junctions in coordinating and regulating the rapid diuretic effects of neuropeptides. Gen. Comp. Endocrinol. 203: 43–48

Fields of Study

Major Field: Entomology

Table of Contents

Abstract ...... ii

Acknowledgments...... iv

Vita ...... v

List of Tables ...... xiv

List of Figures ...... xv

Chapter 1: Introduction ...... 1

Abstract ...... 1

1. Introduction ...... 2

1.1 Control of mosquito borne disease ...... 2

1.2 A brief history of gap junctions ...... 3

1.3 A general comparison of connexins and innexins ...... 5

1.4 Molecular and functional properties of innexins ...... 8

2. Physiological Roles of Innexins ...... 12

2.1 Embryonic development ...... 12

2.2 Nervous system...... 13

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2.3 Reproductive system...... 14

2.4 Immune response ...... 15

2.5 Excretory system ...... 16

3. Goal and rationale of dissertation...... 17

Chapter 2: Pharmacological and genetic evidence for gap junctions as potential new insecticide targets in the yellow fever mosquito, Aedes aegypti ...... 20

Abstract ...... 20

1. Introduction ...... 21

2. Methods ...... 24

2.1 Mosquitoes...... 24

2.2 Chemicals ...... 25

2.3 Adult hemolymph injection assays ...... 25

2.4 Adult topical assays ...... 26

2.5 Adult excretion assay ...... 26

2.6 Larval assays...... 27

2.7 dsRNA synthesis and injection ...... 28

2.8 Phenotype assessment and qPCR ...... 30

2.9 Data analysis and statistics ...... 31

3. Results ...... 32

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3.1 Adult hemolymph injections ...... 32

3.2 Adult topical assays ...... 34

3.3 Adult excretion assays ...... 35

3.4 Larval assays...... 37

3.5 RNA interference (RNAi) ...... 39

4. Discussion ...... 42

4.1 Potential for gap junction inhibitors as insecticides ...... 45

4.2 Conclusions ...... 49

5. Acknowledgements ...... 50

Chapter 3: The molecular and immunochemical expression of innexins in the yellow fever mosquito, Aedes aegypti: insights into putative life stage- and tissue-specific functions of gap junctions ...... 51

Abstract ...... 51

1. Introduction ...... 52

2. Materials and Methods ...... 54

2.1 Mosquitoes...... 54

2.2 Dissections: ...... 54

2.3 Qualitative RT-PCR ...... 55

2.4 Cloning ...... 58

2.5 Sequence Alignment ...... 60

2.6 Anti-Inx3 Antibody ...... 61

2.7 Western Blotting ...... 61

3. Results ...... 64

3.1 Patterns of AeInx expression: ...... 64

3.2 AeInx cDNAs and Gene Structures ...... 68

3.3 Predicted Innexin Proteins: ...... 72

3.4 Immunochemical expression of AeInx3 ...... 75

4. Discussion ...... 80

4.1 General trends ...... 80

4.2 A potential role of AeInx4 in reproduction ...... 81

4.3 A potential role of AeInx8 in development and neuromuscular function: ...... 82

4.4 A potential role of AeInx7 in epithelial and neural function ...... 83

4.5 The molecular potential for homomeric/heteromeric hemichannels and

homotypic/ heterotypic gap junctions ...... 84

4.6 Cloned innexin cDNAs ...... 85

4.7 AeInx3 localization ...... 86

4.8 Summary ...... 88

5. Acknowledgements ...... 88

Chapter 4: Gap junctions and the mosquito blood meal ...... 89

Abstract ...... 89

1. Introduction ...... 90

2. Methods ...... 92

2.1 Mosquito rearing...... 92

2.2 Blood feeding ...... 92

2.3 Dissection, RNA extraction and cDNA synthesis ...... 93

2.4 qPCR ...... 93

2.5 dsRNA synthesis and injections ...... 94

2.6 Fecundity and viability assays ...... 96

2.7 Data Analysis ...... 97

3. Results ...... 98

3.1 Gene expression analysis ...... 98

3.2 RNAi: ...... 103

4. Discussion ...... 105

4.1 The molecular expression of innexins in isolated tissues of non-blood fed

mosquitoes ...... 106

4.2 A blood meal influences innexin mRNA expression ...... 107

4.3 Potential roles of inx2 in the physiology of mosquitoes PBM ...... 108

5. Conclusions ...... 110

6. Acknowledgements ...... 110

Chapter 5: The Contractile Nature of the Mosquito Crop ...... 112

Abstract ...... 112

1. Introduction ...... 113

2. Methods...... 116

2.1 Mosquito rearing...... 116

2.2 Crop dissection ...... 116

2.3 In vitro contraction assays ...... 117

2.4 RNA isolation and qPCR ...... 118

2.5 Antibodies ...... 119

2.6 Immunohistochemistry (IHC) ...... 119

2.7 Statistical analysis...... 120

3. Results ...... 121

3.1 The spontaneous contractions of isolated crops are stable and Ca2+-dependent 121

3.2 Effects of serotonin, benzethonium chloride, and aedeskinin III on crop

contraction rates ...... 123

3.3 Pharmacological and molecular evidence for gap junctions in the crop ...... 124

4. Discussion ...... 128

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4.1 Mechanisms of crop contraction in mosquitoes and other dipterans ...... 128

4.2 Gap junctions and crop contraction: ...... 131

5. Summary and Hypothetical model of crop muscle function ...... 133

6. Acknowledgements ...... 136

Chapter 6: Synthesis ...... 137

References ...... 145

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List of Tables

Table 1: dsRNA template synthesis primers...... 29

Table 2: qPCR primer pairs...... 31

Table 3: Molecular properties of the gap junction inhibitors (rows 2-4) as compared to the properties identified by Tice (2001) for screening for novel insecticides (row 5)...... 48

Table 4: RT-PCR primers used to assess the expression of innexin genes in A. aegypti.

Accession numbers are from Vectorbase (www.vectorbase.org)...... 56

Table 5: Gene specific primers used for RACE...... 59

Table 6: Genbank accession numbers for cloned innexin cDNAs and their splice variants.

...... 60

Table 7: Percent amino-acid identity of the cloned innexins and splice variants to each other and their closest ortholog in D. melanogaster...... 73

Table 8: Phosphorylation sites as identified by PROSCAN analysis...... 75

Table 9: qPCR primer pairs...... 94

Table 10: dsRNA template synthesis primers...... 95

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List of Figures

Figure 1: Structure of gap junctions...... 6

Figure 2: Phylogenetic tree of innexin proteins of Aedes aegypti ...... 10

Figure 3: Dose-response curves of gap junction inhibitors injected directly into the hemolymph of adult female A. aegypti mosquitoes...... 33

Figure 4: Dose-response curves of gap junction inhibitors applied directly to the cuticle of adult female A. aegypti mosquitoes ...... 35

Figure 5: Effects of gap junction inhibitors on the diuretic capacity of adult female A. aegypti mosquitoes...... 37

Figure 6: Dose-response curves of gap junction inhibitors added to the rearing water of 1st instar larval A. aegypti mosquitoes...... 38

Figure 7: Relative innexin gene expression as normalized to RPS7 gene expression in eGFP dsRNA injected mosquitoes 3 days after injection...... 39

Figure 8: Knockdown efficiency in innexin dsRNA injected mosquitoes 3 days after dsRNA injection...... 41

Figure 9: Effect of eGFP (black squares) and innexin (red circles) dsRNA injection on mosquito survival...... 42

Figure 10: Molecular structures of the gap junction inhibitors: A) carbenoxolone, B) mefloquine, and C) meclofenamic acid...... 47

Figure 11: Kyte-Doolittle mean hydrophobicity plot for innexins 1, 2, 3 and 7...... 61

Figure 12: Histogram summarizing the qualitative expression of each innexin in mosquito life stages and sexes...... 65

Figure 13: Histogram summarizing the qualitative expression of each innexin in tissues of the alimentary canal of adult female and male mosquitoes...... 66

Figure 14: Histogram summarizing the qualitative expression of each innexin in the head, carcass, and gonads...... 67

Figure 15: Mapping of the AeInx1, AeInx2, AeInx3, and AeInx7 genes (A, B, C, and D respectively)...... 69

Figure 16: Alignments of the deduced amino-acid sequences encoded by the AeInx1 and

AeInx3 mRNA splice variants...... 70

Figure 17: Amino-acid sequence alignment of the four innexins cloned from adult female

Malpighian tubules...... 73

Figure 18: Western blot of AeInx3 immunoreactivity in a crude lysate of adult female mosquitoes...... 76

Figure 19: Localization of AeInx3-immunoreactivity (green) in isolated tissues of the adult female alimentary canal...... 77

Figure 20: AeInx3 localization...... 79

Figure 21: Knockdown of inx2 in blood fed and non-blood fed mosquitoes...... 96

Figure 22: Tissue specific expression of innexins in non-blood fed females...... 99

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Figure 23: Effects of a blood meal on innexin mRNA expression 3-h post-blood meal. 101

Figure 24: Effects of a blood meal on innexin mRNA expression 24 h post-blood meal.

...... 102

Figure 25: RNAi-induced knockdown of inx2 mRNA expression...... 103

Figure 26: Effects of inx2 knockdown on fecundity of adult female mosquitoes...... 105

Figure 27: In vitro crop assay panel...... 122

Figure 28: Innexin mRNA relative expression levels in the Aedes aegypti crop...... 125

Figure 29: Inx2 and inx3 localization in the crop (ventral diverticulum) of adult female

Aedes aegypti mosquitoes...... 127

Figure 30: Hypothetical model of crop muscle function...... 135

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Chapter 1: Introduction

Abstract

Mosquitoes are the most dangerous animals on the planet due to the pathogens they transmit to humans. For many mosquito borne diseases we do not currently have effective vaccines or therapeutics and instead must control mosquitoes to prevent spread of the diseases. However, mosquitoes are developing resistance to insecticides (e.g., pyrethroids), making the development of new insecticides a necessity. Here we assess the potential of intercellular channels, known as gap junctions, as new molecular targets for insecticide development. Gap junctions allow for direct communication between adjacent cells. All animals possess gap junctions, but they are comprised of evolutionarily distinct families of proteins in vertebrates (connexins) and invertebrates (innexins). Both vertebrate and invertebrate gap junctions play key roles in a diverse range of physiological functions ranging from embryogenesis to reproduction. In Drosophila melanogaster, gap junctions are integral to development, allow for cell coupling in electrical synapses, and are involved in functional gonad formation and gamete production. Recent work by our group in mosquitoes has characterized the molecular expression and immunolocalization

1 of certain innexins and shown that gap junctions are critical in larval and adult mosquitoes. Thus, gap junctions have the potential to be exploited as new targets for chemical- and/or double-stranded RNA-based insecticides.

1. Introduction

1.1 Control of mosquito borne disease

There are over 3,500 species of mosquitoes around the world, which are found on every continent except Antarctica (Harbach 2013). Only a handful of these species represent a threat to human or animal health, yet they are considered the most dangerous animals on Earth (Tolle 2009). Mosquitoes of several genera (Anopheles,

Aedes, and Culex) transmit parasitic and/or viral pathogens, including those that cause malaria, West Nile, dengue fever, and Zika in human hosts. The burden of these diseases is severe, with malaria alone causing over a half million deaths every year, dengue afflicting hundreds of millions of people annually, and Zika being linked to serious birth defects when infecting pregnant women (Bhatt et al. 2013; Shao et al. 2016; Tang et al.

2016; WHO 2015). Moreover, the burden of mosquito-borne diseases extends into animal health, with mosquitoes transmitting pathogens that cause heartworm in canines, eastern equine encephalitis and West Nile virus in horses, and Rift Valley fever in livestock. Unfortunately, many of these diseases lack vaccines and therapeutics; thus,

2 control of the mosquito vectors is typically the primary strategy to control the spread of the diseases.

Mosquito control often relies on the use of insecticides that target the nervous system, such as pyrethroids that modulate voltage-gated sodium channels and carbamates or organophosphates that inhibit acetylcholine esterases. Although these compounds are highly effective at killing mosquitoes, the overuse of a limited diversity of active compounds has exerted a strong selective pressure for the evolution of resistance in the form of target-site resistance (e.g., point mutations in voltage-gated Na+-channels) and/or metabolic resistance. (e.g., up-regulation of cytochrome P450 detoxification enzymes) (Müller et al. 2008; Vontas et al. 2012). Thus, to replenish and diversify our chemical arsenal for controlling mosquitoes, it is necessary to modify existing compounds to by-pass and/or block resistance mechanisms, and/or identify new molecular and physiological targets for developing new insecticides with novel mechanisms of action.

Here, we explore the latter possibility by reviewing the molecular biology and physiology of insect gap junctions within the context of exploiting them as potential targets to develop insecticides for mosquito control.

1.2 A brief history of gap junctions

Gap junctions are intercellular channels that mediate direct communication between adjacent cells via the transfer of small molecules and/or ions. They have been identified in nearly all animals from hydra to humans (Hand and Gobel 1972; Kumar and

Gilula 1986; Takaku et al. 2014). In vertebrates, ultrastructural evidence for gap junctions—hexagonal arrays of proteins in adjacent cell membranes—was first observed in the goldfish (Carassius auratus) brain in what was described as an electrical synapse

(Robertson 1963). It was not until similar structures were discovered in the heart and liver cells of mice that these formations were recognized as new intercellular junctions and the term ‘gap junction’ was coined (Revel and Karnovsky 1967; Revel, Olson, and Karnovsky

1967). In the late 1980s, the genes encoding vertebrate gap junctional subunits, named connexins, were first described in humans and rats (G Dahl et al. 1987; Kumar and Gilula

1986).

In invertebrates, ultrastructural evidence of gap junctions and their subsequent electrophysiological verification were first described in the crayfish (Astacus fluviatilis;

Furshpan and Potter, 1958; Robertson, 1955). Despite this evidence, immunochemical attempts to identify connexin-like molecules in crayfish failed (Berdan and Gilula 1988).

Furthermore, the genomes of both Drosophila melanogaster and Caenorhabditis elegans yielded no homologous connexin genes, indicating that a separate gene family must be responsible for encoding gap junctional subunits in invertebrates (Equence et al. 1998;

Zhao et al. 2000). In the late 1990’s, a family of genes unrelated to the connexins was discovered in D. melanogaster and C. elegans, and demonstrated to form gap junctions in vitro when expressed heterologously in paired Xenopus oocytes; these genes were named ‘innexins’ (invertebrate connexins; Phelan et al., 1998, 1996; Starich et al., 1996).

Intriguingly, vertebrate genomes possess innexin homologues known as pannexins (Baranova et al. 2004). However, pannexins only appear to form hemichannels in the plasma membrane of vertebrate cells and not gap junctions between cells.

Although not involved with direct intercellular communication, pannexins play important functional roles in vertebrate cells, such as mediating the transport of small molecules across the plasma membrane (Gerhard Dahl and Locovei 2006; Scemes and Spray 2012).

Physiological evidence suggests that some innexins are also able to form functional hemichannels in the plasma membrane (L. Bao et al. 2007; Luo and Turnbull 2011). In addition, a viral-based group of innexins (vinnexins) has recently been identified in polydnaviruses. The vinnexins are homologous to insect innexins and can form gap junctions when expressed heterologously in Xenopus oocytes (Turnbull et al 2005). Thus the innexin gene family also includes pannexins and vinnexins, but the primary focus of this review will be insect innexins that form gap junctions.

1.3 A general comparison of connexins and innexins

Connexins and innexins are evolutionarily distinct protein families that have convergently evolved to form gap junctions. The monomeric subunits encoded by the genes share a similar membrane topology that includes four transmembrane domains flanked by intracellular NH2- and COOH- termini (Figure 1A). The transmembrane domains are the most highly conserved and help to form the pore, the extracellular loops are less conserved and are involved in docking, and the termini are the least conserved and

5 involved in gating (Abascal and Zardoya 2013; Bauer et al. 2005). The monomeric subunits form oligomeric hemichannels (typically hexamers) in the plasma membrane, which consist of identical (homotypic) or different (heterotypic) connexin/innexin subunits.

Each hemichannel docks with a complementary hemichannel in the plasma membrane of an adjacent cell to form a pore between the cells, thereby allowing for direct intercellular communication (Figure 1B). The extracellular loops of connexins and innexins possess cysteine residues (Figure 1A), which are thought to mediate the docking of opposing hemichannels (X. Bao et al. 2004). If the molecular composition of the docked hemichannels is identical then the resulting gap junctions are considered homotypic, but if they are different then they are considered heterotypic.

Figure 1: Structure of gap junctions. A) Illustration of the membrane topology of an innexin/connexin gap junctional subunit. Green cysteine residues are found only in connexins, while both innexins and connexins possess the purple residues. B) Illustration of how innexin or connexin subunits combine to form hemichannels and gap junctions, allowing for intercellular communication.

In addition to their similar topology and structure, gap junctions formed by connexins and innexins are similarly gated by intracellular pH, with a rapid decrease leading to channel closure (Camillo Peracchia 2004; Rose, Socolar, and Obaid 1984). Rapid increases in Ca2+ also lead to channel closure (Camillo Peracchia 1978; Rose, Socolar, and

Obaid 1984), potentially via calmodulin binding (Camillo Peracchia et al. 1996). Likewise, most gap junctions are similarly gated by membrane voltage and junctional voltage conductance, with depolarization of the membrane, or increased junctional conductance both leading to increased junctional closure (Bukauskas and Weingart 1994). The subunit composition (i.e. the innexins/connexins forming the gap junction) influence the gating properties that changes in membrane voltage or junctional conductance have on the opening or closure of the gap junctional channels (Alexopoulos et al. 2004; Marks and

Skerrett 2014). Moreover, the open/closed state of gap junctions may be regulated by cell signaling factors. For example, intracellular cAMP is known to increase junctional conductance, presumably through activation of protein kinase A and the phosphorylation of amino acid residues on the NH2 or COOH termini of connexins/innexins (Hax, van

Venrooij, and Vossenberg 1974; De Mello 1983). Additionally, cAMP induced phosphorylation in connexins can stimulate trafficking to the membrane and gap junction formation (Paulson et al. 2000). Thus, sequence variation in the NH2 and COOH termini of

7 connexins/innexins can influence the response of the channels to intracellular factors

(Bauer et al. 2005; Zhang et al. 1999).

In general, the physiological roles of connexins and innexins are remarkably similar. Both are required during embryonic development for proper tissue formation and cell migration (Giuliani et al. 2013; Lo 2000; Ostrowski, Bauer, and Hoch 2008; Rhee et al.

2009), as well as for post-embryonic tissue development (Holcroft et al. 2013; Watanabe and Kankel 1990; Wingard and Zhao 2015). Moreover, connexins and innexins are critical to the functions of excitable and epithelial tissues (Degroot et al. 2003; Furshpan and

Potter 1958; Goliger and Paul 1995; Weng et al. 2008). Below, we focus on the specific molecular and functional properties of innexins, as well as their physiological roles in insects.

1.4 Molecular and functional properties of innexins

In dipteran insects, there are 6 phylogenetic clades of innexins: inx1, inx2, inx3, inx4, inx7, and inx8 (Figure 2). Drosophila melanogaster, mosquitoes, and tsetse flies

(Glossina austeni) each possess a single gene representative of each clade, with the exception of the Inx4 clade, which has diversified via apparent gene duplications to form inx5 and inx6 in D. melanogaster and inx5 in G. austeni (Figure 2). Mosquito genomes typically possess six innexins, which are named after their homologues in D. melanogaster: inx1, inx2, inx3, inx4, inx7, and inx8. Thus, all dipterans appear to have evolved with a core set of at least 6 innexins. The molecular diversity of insect innexins

8 can also be embellished by alternative splicing of mRNA transcripts. At least one innexin in D. melanogaster, inx8, is demonstrated to have at least 3 splice variants, which show differential expression in the giant fiber system (Palacios-Prado, Huetteroth, and Pereda

2014).

Figure 2: Phylogenetic tree of innexin proteins of Aedes aegypti (AaInx), Anopheles gambiae (AgInx), Culex quinquefasciatus (CqInx), Drosophila melanogaster (DmInx), Glossina austeni (GaInx), and one human connexin. (Continued on next page) 10

Figure 2 Continued: The human connexin does not align with any of the innexin proteins as expected. All other innexin proteins sort into the 6 families of Aedes aegypti innexin proteins (inx1, inx2, inx3, inx4, inx7, inx8). Notably, Drosophila melanogaster inx5 and inx6 appear to be duplications that arose after the mosquito and fruit fly ancestors diverged. While there is no Culex quinquefasciatus inx1 in this figure, and none annotated in the genome, a blast of Aedes aegypti inx1 against the Culex genome reveals high homology in an unannotated region (5 potential exons located on supercontig3.776 from 71,153 to 131,352 bp).

To date, the functional properties of specific insect innexins are limited to a few genes in D. melanogaster. Inx8 of D. melanogaster (also known as Shaking-B, ShakB, and passover) was the first insect innexin shown to form functional homotypic gap junctions.

That is, when expressed heterologously in Xenopus oocytes, inx8 mediated the transport of electrical current between paired cells (Pauline Phelan et al. 1998). Moreover, two of the aforementioned differential splice forms of inx8 in D. melanogaster, which differ in the amino-acid sequence of the cytosolic NH2-terminal domain, can form heterotypic gap junctions with novel gating and rectification properties when expressed heterologously in Xenopus oocytes (Phelan et al 2008; Marks and Skerret 2014). Thus, alternative splicing can influence the molecular and functional diversity of the gap junctions formed by innexins.

In contrast to inx8, inx2 and inx3 of D. melanogaster do not consistently form functional homotypic gap junctions when expressed individually in Xenopus oocytes.

Instead, they form functional gap junctions that mediate the transport of electrical current between cells when coexpressed in the same cells (Stebbings et al. 2000). Thus, 11 inx2 and inx3 monomers form heteromeric hemichannels that form homotypic gap junctions.

Specific functional properties of the other innexin members remain to be elucidated, but as described in the following section, molecular expression, reverse genetic, and immunochemical studies, primarily in D. melanogaster, have revealed insights into the broader physiological roles innexins are likely playing in insects.

2. Physiological Roles of Innexins

2.1 Embryonic development

Throughout embryogenesis flies go from a single cell to a multicellular organism with differentiated cells and tissues. This process involves coordination of cells throughout the developing embryo for proper tissue differentiation, orientation, and development. As such, gap junctional communication between developing cells is expected to be critical for embryogenesis. Consistent with this notion, in situ hybridization studies indicate that most innexins undergo differential expression during early embryonic development (Stebbings et al. 2002). Moreover, reverse genetic studies show that the RNAi-driven depletion of inx3 mRNA in D. melanogaster leads to failure to complete dorsal closure in the developing insect (Giuliani et al. 2013).

Innexins are also integral to tissue development. In the central nervous system

(CNS; giant fiber system and ganglia) of D. melanogaster, expression of inx8 isoforms are

12 up and down regulated throughout embryonic development (Zhang et al. 1999).

Moreover, mutations of the inx8 locus lead to impaired CNS development in flies

(Crompton et al. 1995). Innexins are also important in the development of peripheral nerves. Notably, loss of functional inx1 expression, via mutation in the gene locus, results in a severely impaired optical nerve development, inspiring its original name ogre (optical ganglion reduced; Lipshitz and Kankel, 1985; Watanabe and Kankel, 1990). Furthermore,

RNAi mediated knockdown of inx7 results in a severe reduction of peripheral nervous system development (Ostrowski, Bauer, and Hoch 2008).

2.2 Nervous system

Beyond embryonic development of the nervous system, innexins remain integral during larval development. Larvae of D. melanogaster require inx1 for proper cell proliferation in the central nervous system (Pauline Phelan 2005). Inx1 along with inx2 are also required in glial cells for normal neuronal development into adulthood (Holcroft et al. 2013). Moreover, inx1 and inx8 are required for pre- and post- synaptic development of photoreceptor neurons during pupation and for proper neuronal development (Curtin, Zhang, and Wyman 2002). Thus, innexins are needed throughout fly development from embryo to adult.

As adults, D. melanogaster require gap junctions for functional electrical synapses.

Like the giant motor neuron of the crayfish and its role in the tail flip escape response

(Furshpan and Potter 1958; C. Peracchia 1973); D. melanogaster possesses a giant fiber

13 system (GFS) that utilizes electrical synapses for its jump-flight escape response (Allen et al. 2006). In the GFS, mutations leading to non-functional inx8 result in abolishment of electrical synapses in the adult fly and loss of jump response (Phelan et al. 1996).

Moreover, the splice variants of inx8 are differentially expressed throughout the GFS and the splice variants are differentially voltage gated, together allowing for the unidirectional transport of electrical current necessary for electrical synapses (Marks and Skerrett 2014;

Zhang et al. 1999). Innexin-mediated electrical synapses are also involved in connection of the Johnston’s organ’s (fly hearing organ) neuronal circuitry to the GFS, with inx8 and additional unidentified innexin(s) forming these gap junctions (A. Pézier et al. 2014; A. P.

Pézier et al. 2016). Thus, innexins, especially inx8, are critical to proper functioning of the fly nervous system.

2.3 Reproductive system

For both male and female D. melanogaster, inx4 (also known as zero population growth or zpg) is necessary for gonad development and the proper differentiation and survival of developing gametes (Gilboa et al. 2003; Tazuke et al. 2002). Inx4 is also essential to

Anopheles gambiae gonad development (Magnusson et al. 2011). Moreover, RNAi induced gene silencing of inx4 in adult male A. gambiae results in a sterile phenotype that still mates and elicits females to lay non-viable eggs (Thailayil et al. 2011). Inx4 mRNA expression and protein localization occurs in the germline cells of the gonads, which are adjacent to somatic cells that express inx2. This potentially allows for the formation of

14 heterotypic gap junctions between somatic and germline cells, which may be necessary for signaling involved with oogenesis and spermatogenesis (Bohrmann and Zimmermann

2008; Kirilly and Xie 2007; Smendziuk et al. 2015). Inx2 also co-localizes with inx3 in the intercellular membranes of somatic cells in the ovaries, where it is considered necessary for formation of the egg chamber (Bohrmann and Zimmermann 2008; Mukai et al. 2011).

Furthermore, gap junctions mediate coupling between follicle cells and the developing oocyte, allowing the follicle cells to provision the developing oocyte properly. Calmodulin is one such molecule required by the developing oocyte for induction of vitellogen (major yolk protein) uptake (Anderson and Woodruff 2001; Brooks and Woodruff 2004). In D. melanogaster, calmodulin is transferred from the follicle cells to the oocyte via gap junctions (Brubaker-Purkey and Woodruff, 2013). Additionally, in mosquitoes, gap junctions between the oocyte and follicle cells have been observed (Raikhel and Lea

1991). Thus, existing evidence suggests that gap junctions are integral in fly reproduction from gamete production through egg provisioning.

2.4 Immune response

In the insect immune system, gap junctions play important roles in response to invading pathogens and parasites. Electron microscopy studies have shown that gap junctions form between hemocytes during the encapsulation responses, which are part of the innate immune response of insects, in the cockroach (Periplaneta americana) and moth (Calpodes ethlius) (Baerwald 1975; Churchill et al. 1993). In the moth, Spodoptera

15 litura, inx2 and inx3 are expressed in the hemocytes (T. Liu et al. 2013), and may be contributing to this gap junctional formation during encapsulation. D. melanogaster hemocytes similarly encapsulate invading bacteria and parasites, however gap junctions have not yet been implicated in this response (Vlisidou and Wood 2015). Interestingly, polydnaviruses associated with parasitic wasps possess vinnexins (the viral orthologs of innexins) which may alter lepidopteran host gap junction formation by interacting with inx2, thereby disrupting the encapsulation response and allowing the wasp larvae to successfully parasitize its lepidopteran host (Hasegawa and Turnbull 2014; Marziano et al. 2011; Turnbull et al. 2005).

In the mosquito, Anopheles gambiae, inx1 is integral in the up-regulation of

Thioester-containing protein 1 (TEP1), which is a key component of the innate immune response of mosquitoes, after ingestion of blood meals containing malarial parasites

(Plasmodium falciparum) (Li et al. 2014). As such, knockdown of inx1 mRNA expression via RNAi leads to increased invasion and infection of the mosquito by malarial parasites

(Li et al. 2014). Thus, innexins could potentially play an important role in modulating the vector competence of mosquitoes and other disease vectors.

2.5 Excretory system

The renal (Malpighian) tubules of mosquitoes are integral in diuresis after engorging on a blood meal. In the Malpighian tubules of the mosquito, Aedes aegypti, gap junctions are present between the principal cells and allow for electrical coupling

16 between cells as many as five cells apart (Weng et al. 2008). Furthermore, gap junctions may be involved with intercellular signaling required for coordinating the rapid responses of the Malpighian tubule epithelium to diuretic neuropeptides. That is, the receptors for kinin and calcitonin-like peptides, which mediate diuresis, do not occur in every cell along the length of the Malpighian tubules. However, upon treatment with either of these hormones, the epithelium generates a rapid, coordinated physiological response, suggesting robust intercellular communication mediated by gap junctions (Piermarini and

Calkins 2014).

3. Goal and rationale of dissertation

As reviewed above, gap junctions formed by innexins play integral physiological roles throughout the life cycle of insects, especially in dipterans, and numerous additional roles undoubtedly remain to be elucidated. Thus, I hypothesize that gap junctions could serve as valuable targets for the development of new insecticides for mosquito control, because disrupting gap junctions via chemical or genetic inhibition could potentially lead to impaired development, nerve function, reproduction, immune function, and excretion in mosquitoes. Moreover, as indicated earlier, innexins evolved independently of connexins, suggesting that if specific chemical inhibitors of innexins were developed that they could potentially have nominal effects on humans and other vertebrates.

The goal of this dissertation is to further explore the molecular biology and physiological roles of gap junctions in mosquito physiology and evaluate their potential as new insecticidal targets. All of my studies were performed on the yellow fever mosquito, A. aegypti, which is the primary vector of chikungunya, dengue, yellow fever, and Zika viruses. A. aegypti is native to Africa, but now, due to successful invasions, is found in tropical and subtropical regions worldwide. In these studies, I use the Liverpool strain of A. aegypti, which is the strain used to sequence the genome (Nene et al. 2007).

The subsequent chapters document my efforts achieve the goal of my dissertation. In Chapter 2, I use pharmacological and reverse genetic tools to assess whether inhibition of gap junctions is toxic to larval and adult female mosquitoes and disrupts excretory function. In Chapter 3, I use molecular and immunochemical tools to clone innexin cDNAs expressed in the Malpighian tubules, characterize innexin mRNA expression throughout the mosquito life cycle and across major tissues, and localize expression of the Inx3 protein. In Chapter 4, I use molecular tools to assess the effects of a blood meal on innexin mRNA expression in adult females, and explore the consequences of inx2 mRNA knockdown via RNA interference on the survival and fecundity of adult females after a blood meal. In Chapter 5, I evaluate the roles of gap junctions in the muscular contractions mediated by the ventral diverticulum (crop) of adult females using molecular and immunochemical tools along with in vitro physiological assays. In Chapter

6, I summarize the major findings from these chapters and describe their significance in

18 the context of mosquito physiology and the development of novel insecticides for mosquito control.

Chapter 2: Pharmacological and genetic evidence for gap junctions as potential new insecticide targets in the yellow fever mosquito, Aedes aegypti

Published in: Calkins, T.L. and P.M. Piermarini. 2015. Pharmacological and genetic evidence for gap junctions as potential new insecticide targets in the yellow fever mosquito, Aedes aegypti. PLoS One. doi: 10.1371/journal.pone.013708

Abstract

The yellow fever mosquito Aedes aegypti is an important vector of viral diseases that impact global health. Insecticides are typically used to manage mosquito populations, but the evolution of insecticide resistance is limiting their effectiveness.

Thus, identifying new molecular and physiological targets in mosquitoes is needed to facilitate insecticide discovery and development. Here we test the hypothesis that gap junctions are valid molecular and physiological targets for new insecticides. Gap junctions are intercellular channels that mediate direct communication between neighboring cells and consist of evolutionarily distinct proteins in vertebrate (connexins) and invertebrate

(innexins) animals. We show that the injection of pharmacological inhibitors of gap junctions (i.e., carbenoxolone, meclofenamic acid, or mefloquine) into the hemolymph of adult female mosquitoes elicits dose-dependent toxic effects, with mefloquine showing

20 the greatest potency. In contrast, when applied topically to the cuticle, carbenoxolone was the only inhibitor to exhibit full efficacy. In vivo urine excretion assays demonstrate that both carbenoxolone and mefloquine inhibit the diuretic output of adult female mosquitoes, suggesting inhibition of excretory functions as part of their mechanism of action. When added to the rearing water of 1st instar larvae, carbenoxolone and meclofenamic acid both elicit dose-dependent toxic effects, with meclofenamic acid showing the greatest potency. Injecting a double-stranded RNA cocktail against innexins into the hemolymph of adult female mosquitoes knock down whole-animal innexin mRNA expression and decreases survival of the mosquitoes. Taken together these data indicate that gap junctions may provide novel molecular and physiological targets for the development of insecticides.

1. Introduction

The yellow fever mosquito, Aedes aegypti, is the most important vector of the viruses that cause yellow, dengue, and chikungunya fevers in humans. These diseases have spread around the tropical and subtropical world, facilitated by globalization of human societies and climate change (Tolle 2009). In particular, chikungunya fever has most recently emerged from its native range in sub-Saharan Africa and Asia to Central

America and the Caribbean in 2013-2014. As of 2014, locally acquired cases of chikungunya were reported in Florida (CDC 2015; Powers 2014).

Ideally, these mosquito-borne diseases could be prevented through the global use of safe, effective, and affordable vaccines. For yellow fever there is such a vaccine, however for chikungunya and dengue fevers there currently are no effective vaccines available (WHO 2014). An alternative strategy for controlling the spread of mosquito- borne diseases is to control populations of the mosquito vectors that transmit the pathogens. The primary control methods for reducing mosquito numbers are sanitation

(cleaning and removing larval habitats from around homes) and using insecticides (CDC

2013). Although insecticides are effective at reducing mosquito populations, insecticide resistant populations have emerged because of the overuse of a few limited active compounds, such as pyrethroids (Müller et al. 2008). The control of resistant populations of mosquitoes can be mitigated through a variety of techniques, including the development of new insecticides with novel modes of action, which begins with the identification of new insecticidal targets.

Gap junctions are potential molecular and physiological targets for the development of new insecticides. On the cellular level, gap junctions are intercellular channels that allow for the transport of small molecules and ions between adjacent cells

(Cao et al. 1998). On the molecular level, gap junctions are formed by two hemichannels from neighboring cells that dock with one another. Each hemichannel consists of six protein subunits, which are encoded by genes called connexins in vertebrates and innexins in invertebrates. The connexin and innexin proteins possess similar structural and functional features, but have evolved independently and thus their primary

22 structures possess little similarity to one another (P Phelan and Starich 2001; Swenson et al. 1989).

In the A. aegypti genome, 6 genes encode innexins (Weng et al 2008); we have demonstrated that these genes are differentially expressed throughout the mosquito life cycle and in various tissues of adult mosquitoes (Calkins et al. 2015; Weng et al. 2008). In insects, innexins are known to play key roles in embryogenesis. For example, knockout of innexin 3 (inx3) in Drosophila melanogaster results in a failure of dorsal closure (Giuliani et al. 2013). Moreover, in Anopheles gambiae, disruption of innexin 4 (a.k.a ‘zero- population growth’ or ‘zpg’) results in sterile males (Thailayil et al. 2011). Innexins are also thought to be important in adult insect neuromuscular communication and renal function (Anava et al. 2009; Piermarini and Calkins 2014; Weng et al. 2008). In summary, given the evolutionary distinct ancestry of innexins and connexins, as well as the consequences of innexin disruption on insect biology, we hypothesized that gap junctions may serve as valuable targets for insecticide development.

To test our hypothesis, we assessed the effects of gap junction inhibition on mosquito survival and/or physiology using pharmacological and genetic tools. In particular, we used three commercially available gap junction inhibitors (carbenoxolone, meclofenamic acid and mefloquine), which block vertebrate and invertebrate gap junctions formed by connexins and innexins, respectively (Cruikshank et al. 2004; Luo and

Turnbull 2011; Sangaletti, Dahl, and Bianchi 2014; Srinivas and Spray 2003). Furthermore,

23 we utilized RNA interference (RNAi) to knock down the expression of the 6 innexin mRNAs expressed in adult female A. aegypti.

We find that all 3 pharmacological inhibitors are toxic to adult female mosquitoes when injected into the hemolymph, and that carbenoxolone is effective when applied topically to the cuticle. Moreover, carbenoxolone and mefloquine decrease the diuretic capacity of adult female mosquitoes, suggesting disruption of Malpighian tubule function as a potential mechanism of action for these compounds. Also, carbenoxolone and meclofenamic acid kill 1st instar larvae when added to their rearing water. Lastly, the knockdown of innexin mRNA levels in adult female mosquitoes decreases their survival over 11 days. Taken together, our results indicate that gap junctions are promising molecular and physiological targets for the development of novel insecticides to control mosquito vectors.

2. Methods

2.1 Mosquitoes

Eggs of Aedes aegypti were obtained through the Malaria Research and Reference

Reagent Resource Center (MR4) as part of the BEI Resources Repository (Liverpool strain;

LVP-IB12 F19, deposited by M.Q. Benedict). Mosquitoes were reared as described in

Piermarini et al. (2011) in an environmental chamber set at 28C and 80% relative humidity with a 12 h:12 h light:dark cycle.

2.2 Chemicals

Carbenoxolone, meclofenamic acid and mefloquine were obtained from Sigma-

Aldrich (St. Louis, MO). All other chemicals were obtained from Thermo Fisher Scientific

(Waltham, MA).

2.3 Adult hemolymph injection assays

For direct hemolymph injection, carbenoxolone and meclofenamic acid were dissolved in HEPES buffered saline (HBS) as 100 mM stock solutions, whereas mefloquine was dissolved into 100% dimethyl sulfoxide (DMSO). Before injection, the inhibitors were diluted to their desired concentrations in HBS. The HBS consisted of 11.9 mM HEPES, 137 mM NaCl, and 2.7 mM KCl; the pH was adjusted to 7.45 using NaOH. For dilutions of carbenoxolone and meclofenamic acid, DMSO was added to the HBS at a final concentration of 11% to match that found in dilutions of mefloquine.

Adult female mosquitoes (3-10 days post-eclosion) were immobilized on ice prior to injecting their hemolymph with 69 nl of an inhibitor using a Nanoject II microinjector

(Drummond Scientific Company, Broomall, PA). For a given dose of a compound, ten mosquitoes were injected and transferred to small cages (10 mosquitoes per cage) with access to a 10% sucrose solution. The cages were returned to the rearing chamber and the efficacy of a dose was assessed 24 h later, as described in Raphemot et al. (2013). In brief, the efficacy was measured as the percentage of treated mosquitoes in a cage that

25 were incapacitated by 24 h; i.e., the collective percentage of mosquitoes that were flightless or dead (Raphemot et al 2013). A total of five to ten independent replicates were performed for each dose of each inhibitor.

2.4 Adult topical assays

For topical application, all inhibitors were dissolved directly at their desired concentrations in 75% ethanol/25% H2O. Adult female mosquitoes (3-10 days post- eclosion) were immobilized on ice and a Hamilton repeating dispenser (Hamilton

Company, Reno, Nevada) was used to apply 500 nl of an inhibitor to the thorax of each mosquito. For a given dose of compound, ten mosquitoes were treated and transferred to small cages (10 mosquitoes per cage) with access to a 10% sucrose solution. The cages were returned to the rearing chamber and the efficacy of a dose was assessed after 24 h, as described above in ‘Adult hemolymph injection assays’. A total of four to eight independent replicates were performed for each dose of each inhibitor.

2.5 Adult excretion assay

The diuretic capacity of adult female mosquitoes was assessed using an established protocol (Raphemot et al. 2013). In brief, for a given treatment, 3 mosquitoes

(3-10 days post eclosion) were immobilized on ice and injected with 900 nl of HBS into their hemolymph using a Nanoject II microinjector (Drummond Scientific). After injection, the mosquitoes were placed into a graduated, packed-cell volume tube (MidSci, St. Louis,

MO) for two hours at 28 C to allow them to excrete. After two hours, the mosquitoes were removed from the tube, which was then centrifuged at 17,000 g to allow for the excreted volume to be measured visually via the graduated column at the bottom of the tube. At minimum, 6 replicates (3 mosquitoes per replicate) were performed for each treatment. All mosquitoes were confirmed to be alive at the end of the two hours.

Mosquitoes that were not injected with HBS served as controls.

The composition of the HBS was similar to that described above in ‘Adult hemolymph injection assays’ except that NaOH and DMSO were reduced to 1 mM and

2%, respectively. For a given experiment, one of the following inhibitors was added to the HBS at the indicated concentrations: carbenoxolone (1.34 mM), mefloquine (0.5 mM), and meclofenamic acid (1.53 mM).

2.6 Larval assays

The toxicity of the inhibitors on 1st instar larvae was assessed using the protocol of Pridgeon et al. (2009). In brief, eggs were hatched in dH2O under vacuum at room temperature for 2 h in a 250 ml beaker. The beaker was transferred to a rearing chamber

(28°C) and 1st instar larvae were collected 24 h later. For a given treatment, 5 larvae were transferred to the well of a 24 well Falcon MULTIWELL plate (Becton Dickinson Labware,

Franklin Lakes, NJ) containing 995 µl of H2O with a gap junction inhibitor and 5 µl of food solution. The food solution consisted of 0.013 g of ground Tetramin fish food (Tetra

United Pet Group, Blacksburg, VA) suspended in 1 ml of dH2O. Stock solutions of the

27 inhibitors were dissolved in dH2O and diluted within wells to achieve the desired concentrations (1 ml total volume per well). Plates were returned to rearing conditions and after 24 hours the larvae were assessed. The efficacy of a concentration was measured as the percentage of treated larvae in a well that were dead by 24 h. Larvae were considered dead if they did not move when disturbed with a 10 µl pipette tip.

2.7 dsRNA synthesis and injection

dsRNAs were synthesized from DNA templates. To generate the DNA templates, primers were designed for each innexin cDNA using Primer3 software (Rozen and

Skaletsky 1998) to amplify a 300-500 base pair product (Table 1). The nucleotide sequences of the primers and expected PCR products were subjected to a Basic Local

Alignment Search Tool (BLAST) against the A. aegypti genome (Vectorbase.org) to ensure specificity. A T7 promoter sequence (TAATACGACTCACTATAGGGAGA) was added to the

5’ end of each forward and reverse primer to allow for dsRNA synthesis. The DNA templates were generated via PCR, which consisted of mixing 0.5 µl plasmid DNA (from previously cloned innexin cDNAs), 1 µl forward and reverse primer mix (10 µM each) and

23.5 µl Platinum® Taq DNA polymerase High Fidelity (Thermo-Fisher). The mixture was subjected to a thermocycling protocol consisting of an initial denaturation at 95 °C (2 min) followed by 30 cycles of 95 °C (1 min), 60 °C (30 sec), 72 °C (1 min); the protocol ended with an elongation step at 72 °C (5 min). The identities of the various DNA templates generated by PCR were verified by agarose gel (1%) electrophoresis and Sanger DNA

28 sequencing at the Molecular and Cellular Imaging Center (MCIC) of the Ohio Agricultural

Research and Development Center (OARDC) of the Ohio State University (Wooster, OH).

Table 1: dsRNA template synthesis primers. Each primer set consists of an innexin specific region for amplification of the target gene from plasmid, and the T7 promoter sequence (TAATACGACTCACTATAGGGAGA).

dsRNA Template Forward dsRNA Template Reverse Inx1 TAATACGACTCACTATAGGGAGAGC TAATACGACTCACTATAGGGAGAAAA GAAGCTGCAGAAGCTATT TGTTTTGTCGAGGTTCATGT Inx2 TAATACGACTCACTATAGGGAGATTT TAATACGACTCACTATAGGGAGAATA GGCGTTTGAAAAGTGTG CTCCCGGCTGAGCAATA Inx3 TAATACGACTCACTATAGGGAGACG TAATACGACTCACTATAGGGAGAGTT ACGGTGACAGATTGACTAG CGCTCCTGGTTGTACTC Inx4 TAATACGACTCACTATAGGGAGACA TAATACGACTCACTATAGGGAGAAA TTCCTGTTCTCGTTCCCC GGCACAGGGCATCAAAGT Inx7 TAATACGACTCACTATAGGGAGACA TAATACGACTCACTATAGGGAGATCA GGGACAATCCAAAAGCATG GTTTCGTCAGCCTCATC Inx8 TAATACGACTCACTATAGGGAGATT TAATACGACTCACTATAGGGAGATCA CTGACGATACTGACGACGTT TGCATCCTGTATTTCACCT eGFP TAATACGACTCACTATAGGGACGTA TAATACGACTCACTATAGGGTTGGGG AACGGCCACAAGTT TCTTTGCTCAGG

Template DNA was then used in the T7 MEGAscript® dsRNA synthesis kit (Thermo-

Fisher Scientific) following the manufacturer’s protocol (20 µl total reaction Volume). The resulting dsRNA was resuspended in nuclease free water and its concentration was measured on a Nanodrop 2000 spectrophotometer (Thermo Scientific). The dsRNA for each innexin was diluted to approximately 4 µg/µl, aliquoted, and stored at -80 °C to avoid repeated freeze-thaw degradation.

On the day of an experiment, all six innexin dsRNAs were diluted to 2 µg/µl in a

PBS solution (137 mM NaCl, 2.7 mM KCl and 11.9 mM phosphates; pH 7.5). Before injection into a mosquito, equal volumes of each innexin dsRNA were mixed together to form an innexin dsRNA ‘cocktail’, resulting in a final concentration of ~333 ng/µl for each dsRNA. Adult female mosquitoes were anesthetized on ice and their hemolymph was injected with either 1 µl of the innexin dsRNA cocktail or a negative control dsRNA against enhanced green fluorescent protein (eGFP; 2 µg/µl) using a Nanoject II injector

(Drummond Scientific). For the eGFP and innexin dsRNAs, a total of 30 mosquitoes were injected per replicate (6 mosquitoes were dedicated for qPCR analysis and 24 were dedicated for survival assays). After injection all mosquitoes were placed in small cages and returned to rearing conditions. Four biological replicates were performed for knockdown analysis and three biological replicates for the survival assay.

2.8 Phenotype assessment and qPCR

Mosquitoes injected with eGFP or innexin dsRNA were checked at 24 h intervals for 11 days and the number of surviving mosquitoes was recorded. Real time quantitative

PCR (qPCR) was utilized to determine knockdown on days 3, 7 and 11 post-injection. RNA extraction and cDNA synthesis were performed as described in Calkins et al. (2015).

Primers for qPCR were designed against the six innexins and ribosomal protein S7 (RPS7; a reference gene) using Primer3 software (Rozen and Skaletsky, 1998) to amplify a 90-110

30 base pair product (Table 2). Specificity of the resulting PCR products was confirmed using a melt curve analysis and Sanger sequencing (MCIC, OARDC).

Table 2: qPCR primer pairs. Each set of primers was selected for innexin specificity and determined specific through melt curve analysis and sequencing (MCIC, OARDC).

qPCR Forward qPCR Reverse Inx1 CACCGATAGTGCCGTATTCC CCGACATATTGTGTGGCAGT Inx2 GGAGATCCTATGGCACGAGT ACGGTAGCACACAGAGTCCA Inx3 TCGTTCGGTTACTTCATCTGC GCGATTCTCCTGATCCATGTC Inx4 TTCTGTTGGACACTGGGAAC CCATGTGCGTTCCTATTTCG Inx7 TGGGTCCCGTTTGTGTTATT CCATACGAAGACCATCCACA Inx8 GACTGCGTTCACACGAAAGA GGGTACTTCGCTACCGACTTT RPS7 CTTTGATGTGCGAGTGAACAC CATCTCCAACTCCAGGATAGC

For a given sample, each qPCR consisted of three technical replicates of 10 µl reactions each consisting of 5 µl of GoTaq® Master Mix, 40 ng cDNA, 400 nM forward and reverse primers, and nuclease free water. The reactions took place in 96-well unskirted, low profile plates (Bio-Rad Laboratories, Hercules, CA), sealed with TempPlate® RT optical film (USA Scientific). qPCR was performed using a Bio-Rad C1000 thermocycler and CFX96 real time system (Bio-Rad Laboratories). The thermocycler used the following protocol: an initial denaturation of 95 °C (3 min) followed by 39 cycles of 95 °C (10 sec) and 58 °C

(30 sec), ending with a melt curve cycle.

2.9 Data analysis and statistics

GraphPad Prism 6 software (GraphPad Software Inc., La Jolla, CA) was used in all statistical analysis. Results of the toxicology experiments with gap junction inhibitors (i.e., adult hemolymph injections, adult topical applications, and larval assays) were analyzed using a non-linear curve fit analysis (log [inhibitor] vs. response variable-slope) to determine effective dose or concentration 50% values (i.e., ED50 or EC50). Results from the excretion assays were analyzed with a one-way ANOVA with a Newman-Keuls post hoc analysis. Relative gene expression was determined utilizing the delta CT method by normalizing target gene expression to that of the reference gene RPS7. Relative gene expression levels in eGFP controls were analyzed with a one-way ANOVA with a Newman-

Keuls post hoc analysis. Percent gene silencing was calculated as in Drake et al. (2010) by setting relative gene expression in eGFP injected mosquitoes to 100%. Significant knockdown was determined via Student’s t-tests comparing normalized innexin mRNA levels in eGFP dsRNA injected mosquitoes to that of innexin dsRNA injected mosquitoes.

Survival between eGFP and innexin injected groups was compared by a two-way repeated measures ANOVA with Holm-Sidak’s post hoc analysis.

3. Results

3.1 Adult hemolymph injections

To determine if gap junction inhibitors are toxic to adult female mosquitoes, we injected the inhibitors directly into the hemolymph. Carbenoxolone, mefloquine and meclofenamic acid all showed dose-dependent toxic effects in adult female mosquitoes

(Figure 3). Mefloquine was the most effective inhibitor, with an effective dose for 50% of the population (ED50) of 15.47 ng per mg of mosquito body weight (ng/mg) followed by meclofenamic acid (ED50 = 96.39 ng/mg) and carbenoxolone (ED50 = 127.3 ng/mg; Figure

1).

Figure 3: Dose-response curves of gap junction inhibitors injected directly into the hemolymph of adult female A. aegypti mosquitoes (carbenoxolone R2=0.873, meclofenamic acid R2=0.957 and mefloquine R2=0.906). Efficacy (dead and flightless mosquitoes) was assessed 24 h after injection. Taking into consideration the average mass of an adult female mosquito (1.97 mg), the ED50 for carbenoxolone, meclofenamic acid and mefloquine are 127.3 ng/mg, 96.4 ng/mg and 15.47 ng/mg respectively. Values are means ± SEM. n=5-10 replicates of ten mosquitoes per dose tested.

3.2 Adult topical assays

To determine whether the gap junction inhibitors can penetrate the cuticle, we evaluated the efficacy of the inhibitors in adult female mosquitoes when applied topically to the thorax. Of the three inhibitors, carbenoxolone was the only one to show a dose- dependent effect nearing 100% efficacy, with an ED50 of 8.57 µg/mg. Mefloquine showed limited dose-dependent effects, with a maximal efficacy of only ~54.5%. Meclofenamic acid showed the weakest topical efficacy (Figure 4).

Figure 4: Dose-response curves of gap junction inhibitors applied directly to the cuticle of adult female A. aegypti mosquitoes (carbenoxolone R2=0.824, meclofenamic acid R2=0.455 and mefloquine R2=0.44). Efficacy (dead and flightless mosquitoes) was assessed 24 h after application. Taking into consideration the average mass of an adult female mosquito (1.97 mg), the ED50 for carbenoxolone is 8.57 µg/mg. The ED50s for meclofenamic acid and mefloquine are not determinable. Values are means ± SEM. n=4- 8 replicates of ten mosquitoes per dose tested.

3.3 Adult excretion assays

To determine if the gap junction inhibitors disrupt the diuretic capacity of adult female mosquitoes, we volume loaded their hemolymph with a sub-lethal dose of each

35 inhibitor. Figure 3 shows the mean volume excreted per female in 2 h after injection for each treatment, compared to the non-injected controls (C). Mosquitoes injected with a volume load (VL) and no inhibitor excreted on average 760 ± 16 nl (Figure 5). In contrast, those injected with a VL and 1.34 mM carbenoxolone (VL + CBX) excreted a significantly lower amount of urine (8.0 ± 8 nl) that is similar to non-injected control mosquitoes, which excrete 39 ± 8 nl (Figure 3). Mosquitoes injected with a VL and 0.5 mM mefloquine (VL +

MEF) excrete 290 ± 93 nl, which was significantly lower than the amount excreted by the

VL mosquitoes, but significantly higher than that excreted by the VL + CBX mosquitoes

(Figure 3). Mosquitoes injected with a VL and 1.53 mM meclofenamic acid (VL + MFA) excrete 844 ± 14 nl, which was comparable to that of VL mosquitoes (Figure 3).

Concentrations of meclofenamic acid higher than 1.53 mM were lethal to the mosquitoes before the end of the 2 h excretion assay (data not shown).

Figure 5: Effects of gap junction inhibitors on the diuretic capacity of adult female A. aegypti mosquitoes. C = non-injected control mosquitoes. VL= volume loaded mosquitoes injected with 900 nl of HBS. VL + CBX = mosquitoes injected with 900 nl of HBS and 1.34 mM carbenoxolone. VL + MEF = mosquitoes injected with 900 nl of HBS and 0.5 mM mefloquine. VL + MFA = mosquitoes injected with 900 nl of HBS and 1.53 mM meclofenamic acid. Letters (A, B or C) indicate statistical differences as determined by a one-way ANOVA and Newman-Keuls post-test (P < 0.05). Values are mean volumes of urine excreted per mosquito after two hours ± SEM. n=14 for VL, n=14 for C, n= 8 for VL + CBX, n=6 for VL + MEF, and n = 6 for VL + MFA.

3.4 Larval assays

We assessed the efficacy of the gap junction inhibitors as larvicides by adding them to the rearing water of 1st instar larvae. Carbenoxolone and meclofenamic acid both

37 showed dose-dependent toxic effects in larvae (Figure 6). Meclofenamic acid was the most effective (EC50 = 244.9 ppm) followed by carbenoxolone (EC50 = 1587 ppm). We were unable to test the efficacy of mefloquine in this assay, because it is not soluble in water and the amount of DMSO required to keep it in solution (>2%) was toxic to larvae.

Figure 6: Dose-response curves of gap junction inhibitors added to the rearing water of 1st instar larval A. aegypti mosquitoes (carbenoxolone R2=0.873 and meclofenamic acid 2 R =0.957). Larval mortality was assessed 24 h after adding the inhibitors. LC50 for meclofenamic acid and carbenoxolone are 0.83 mM and 2.84 mM, respectively. Values are means ± SEM. n=4-8 replicates of five larvae per concentration tested.

3.5 RNA interference (RNAi)

To determine if knockdown of innexin mRNA levels affected the survival of adult female mosquitoes, we utilized RNAi. First, we used qPCR to determine the relative expression of each innexin in mosquitoes injected with eGFP dsRNA (3 days post injection). As shown in Figure 7, Inx2 was the most abundant innexin, followed by Inx3 and Inx 4. The expression of Inx1, Inx7 and Inx8 were lower, but detectable.

Figure 7: Relative innexin gene expression as normalized to RPS7 gene expression in eGFP dsRNA injected mosquitoes 3 days after injection. Values are means ± SEM. n = 4 replicates of 3 mosquitoes. Letters (A, B or C) indicate statistical differences as determined by a one-way ANOVA and Newman-Keuls post-test (P < 0.05).

Injection of an innexin dsRNA cocktail containing dsRNAs for each innexin (333 ng per innexin, 2 µg total) resulted in significant knockdown of Inx1 (32 ± 8%), Inx2 (69 ± 3%),

Inx3 (51 ± 5%), Inx4 (71 ± 10%) and Inx7 (86 ± 2%) by 3 days after injection compared to expression levels in the eGFP-injected controls (Figures 8). The expression levels of Inx8 were not significantly knocked down, but the mRNA levels were very low to begin (Figure

5). The knockdown of innexin expression in mosquitoes injected with innexin dsRNA persisted until at least day 11 (data not shown). In addition, mosquitoes injected with innexin dsRNA exhibited a significantly lower survival than those injected with eGFP dsRNA that progressed over the next 11 days, and started as early as 1 day after injection

(Figure 9).

Figure 8: Knockdown efficiency in innexin dsRNA injected mosquitoes 3 days after dsRNA injection. Percent knockdown is relative to eGFP dsRNA injected control mosquitoes 3 days after injection (Figure 5). Values are means ± SEM. n = 4 replicates of 3 mosquitoes. Asterisks indicate significant knockdown compared to eGFP as determined by a Student’s t-test (p < 0.05).

Figure 9: Effect of eGFP (black squares) and innexin (red circles) dsRNA injection on mosquito survival. Values are means ± SEM. n = 3 replicates of 24 mosquitoes. Asterisks indicate a significant difference in survival between eGFP and innexin dsRNA injected mosquitoes as determined by a two-way ANOVA with a Holm-Sidak’s post-hoc test.

4. Discussion

Our study provides the first pharmacological and molecular evidence that suggests gap junctions are potentially valuable targets for mosquitocide discovery and development. When injected into the hemolymph of adult female mosquitoes, three structurally unique pharmacological agents that are known to inhibit gap junctions

(carbenoxolone, meclofenamic acid, and mefloquine) were efficacious (Figure 3), and one 42 of the compounds (carbenoxolone) exhibited the ability to penetrate the cuticle of adult females (Figure 4). Moreover, two of the gap junction inhibitors (carbenoxolone and meclofenamic acid) were effective when added to the rearing water of 1st instar larvae

(Figure 6). Thus, our data provide proof-of-concept that chemical inhibitors of gap junctions exhibit insecticidal properties in adult and larval mosquitoes.

In support of our pharmacological data, the injection of dsRNA against all 6 innexin mRNAs reduced the survival of adult female mosquitoes over the next 11 days (Figure 9).

The reduced survival was associated with the significant knockdown of mRNA levels for 5 of the targeted innexins (Figure 8), which presumably leads to reductions of innexin protein levels and higher mortality. The mRNA levels of Inx8 were not significantly affected by dsRNA injection, but this gene is expressed at nominal levels in adult female mosquitoes (Figure 7; Calkins et al 2015). Notably, the degree and rate at which the toxic effects manifest via RNAi are respectively weaker and slower than those of the pharmacological inhibitors. The weaker effects of RNAi are likely attributable to a combination of the incomplete knockdown of innexin mRNA expression (Figure 8) and the time required for dsRNAs to elicit an effect on protein levels, which may lag behind that of mRNA levels. In contrast, pharmacological inhibition can be complete and elicit acute effects. Other studies have also noticed weaker toxic effects of inhibition of an insecticide target via RNAi vs. pharmacological inhibition (Raphemot et al. 2014; Revuelta et al. 2009;

Zhou and Xia 2009).

The toxic effects of the gap junction inhibitors and the innexin dsRNAs on mosquitoes are not surprising given that innexin mRNAs are expressed throughout the mosquito life cycle (Calkins et al 2015). Furthermore, in adult female mosquitoes, we have shown that at least 4 innexin mRNAs are expressed in each tissue of the alimentary canal

(i.e., midgut, hindgut and Malpighian tubules), the ovaries, head, and thorax/abdomen

(Calkins et al. 2015). Thus, the pharmacological or genetic inhibition of innexin function may cause disruptions to the nervous, digestive, excretory, and/or reproductive systems, leading to the impairment of flight and/or death. Further investigations will be required to confirm that such wide-spread disruptions to mosquito physiology are indeed occurring.

In the present study, we show that at least two of the pharmacological inhibitors

(carbenoxolone and mefloquine) perturb the functions of the excretory system, as indicated by their inhibition of the diuretic capacity of adult female mosquitoes (Figure

5). These two inhibitors may be acting on the Malpighian tubules, which produce urine via transepithelial fluid secretion, and/or the hindgut, which attenuates the composition of urine before expelling it from the animal via muscular contractions. In Malpighian tubules, several lines of evidence suggest that gap junctions composed of innexins occur between the epithelial cells and play important roles in intercellular communication and diuresis (K W Beyenbach and Piermarini 2011; Lu, Kersch, and Pietrantonio 2011;

Piermarini et al. 2010; Piermarini and Calkins 2014). Furthermore, in the hindgut, we have localized the expression of Inx3 immunoreactivity to the intercellular membranes of

44 epithelial cells in the ileum and rectum (Calkins et al. 2015). Thus, inhibiting the activity of gap junctions in these tissues is expected to disrupt urine production and/or expulsion.

However, additional experiments such as Ramsay assays of isolated Malpighian tubules and isolated hindgut contraction assays (Nachman et al. 1997; Ramsay 1953) are needed to resolve the mechanisms by which carbenoxolone and mefloquine inhibit excretory performance.

Surprisingly, we did not observe effects of a sub-lethal dose of meclofenamic acid

(1.53 mM) on the diuretic capacity of mosquitoes (Figure 5), and higher doses were lethal before the 2 hr experimental period finished (Calkins, unpublished observations). These findings suggest that meclofenamic acid either does not inhibit gap junctions expressed in the excretory system of mosquitoes, or it may elicit rapid toxic effects elsewhere, such as in the nervous system before any effects on excretory function can be observed.

Perhaps, meclofenamic acid is able to cross the blood-brain barrier of mosquitoes more efficiently than carbenoxolone and mefloquine, thereby leading to a more rapid toxic effect.

4.1 Potential for gap junction inhibitors as insecticides

When applied topically to the cuticle of mosquitoes, only carbenoxolone showed insecticidal activity, whereas mefloquine and meclofenamic acid had limited and nominal activity, respectively. Thus, carbenoxolone appears to have the capacity to penetrate the cuticle. The structures (Figure 10) and chemical properties (Table 3) of these three gap

45 junction inhibitors are distinct and may explain their different abilities to penetrate the cuticle. For example, when evaluating the three inhibitors based on the “Rule of 5”

(Lipinski et al. 2001; Tice 2001) for identifying insecticides (Table 3), carbenoxolone adheres closely with a ‘clog p’ below 5, zero hydrogen bond donors, 7 hydrogen bond acceptors, only 6 rotatable bonds, and a molecular weight near 500 kDa (Table 3). Besides molecular weight, the other two gap junction inhibitors differ from carbenoxolone primarily in their number of hydrogen bond donors. Carbenoxolone is the only one of the three compounds that adheres to the norm of current insecticides (Tice 2001) with no hydrogen bond donors.

Figure 10: Molecular structures of the gap junction inhibitors: A) carbenoxolone, B) mefloquine, and C) meclofenamic acid. Structures as shown were modified from Sigma- Aldrich (www.sigma.com) and constructed in ChemSketch (ACD/Labs, Toronto, Ontario, Canada).

Table 3: Molecular properties of the gap junction inhibitors (rows 2-4) as compared to the properties identified by Tice (2001) for screening for novel insecticides (row 5).

Hydrogen Hydrogen Molecular Rotatable Inhibitor cLog P Bond Bond Weight Bonds Donors Acceptors Carbenoxolone 570.77 4.83 0 7 6 Mefloquine 378.32 3.70 2 3 4 Meclofenamic 296.15 4.92 2 3 3 Acid Tice Rule of 5 < 500 < 5 0 < 10 < 12

Despite nominal topical efficacy against adult female mosquitoes, meclofenamic acid was the most potent larvicide of the three compounds (Figure 6). Carbenoxolone was also effective against larvae, but over 6-fold less potent than meclofenamic acid. Given that meclofenamic acid was unable to penetrate the cuticle of adult females (Figure 6), we presume that in larvae this compound is delivered to the alimentary canal via ingestion where it may act on the midgut epithelium and/or diffuse into the hemolymph.

Although meclofenamic acid and carbenoxolone were respectively the most effective compounds against larval and adult female (topically) mosquitoes, it is important to emphasize that these compounds are not nearly as potent as conventional insecticides, such as permethrin. For example, meclofenamic acid is 46 times less effective against 1st instar larvae of A. aegypti than DNOC (Dinitro-ortho-cresol), which is considered a weak insecticide, and 874,643 times less effective than permethrin, a highly potent insecticide (Pridgeon et al. 2009). Additionally, our most topically active inhibitor,

48 carbenoxolone, is 5.7 times less effective than bifenzate, a very weak mosquitocide and over 18.5 million times less effective than fipronil, the most potent adulticide against A. aegypti (Pridgeon et al. 2009).

Additional concerns for both carbenoxolone and meclofenamic acid are their efficacy on mammalian gap junctions (Carlen et al. 2000; Ning et al. 2013). Moreover, carbenoxylone has potential off target effects in mammals, and meclofenamic acid, is currently used therapeutically as a nonsteroidal anti-inflammatory drug (NSAID) pain reliever (Connors 2012; Ning et al. 2013). Thus, our data should only be taken as proof- of-concept that gap junction inhibitors possess insecticidal properties. Additional efforts will be necessary to modify the potency and selectivity of these compounds for mosquitoes before they could be considered for use as insecticides in the field.

4.2 Conclusions

The present study is the first to demonstrate that three different commercially available gap junction inhibitors exhibit insecticidal activity. Moreover, we show that

RNAi-based knockdown of mRNAs encoding gap junctional proteins (i.e., innexins) leads to increased mortality of adult female mosquitoes. Taken together, these results suggest that gap junctions offer new potential insecticidal targets for mosquito control. However, much progress still needs to be made in terms of discovering compounds with acceptable potency and specificity for mosquito gap junctions. The long evolutionary distance

49 between innexins and connexins leaves us hopeful that an innexin-specific compound can be discovered and developed.

5. Acknowledgements

We thank Dr. Matthew Rouhier (Kenyon College) for his assistance in the

Laboratory, and Ms. Nuris Acosta (OSU) and Ms. Edna Alfaro (OSU) for their assistance in mosquito rearing. Funding for this study was provided by grants to 1) PMP from the

NIH (R03DK090186) and Mosquito Research Foundation (2014-03), and 2) TLC from the

OARDC SEEDS program (Grant# 2014-078; oardc.osu.edu/seeds) and Ohio Mosquito

Control Association Grant-In-Aid. State and Federal funds appropriated to the OARDC of the Ohio State University also supported the study.

Chapter 3: The molecular and immunochemical expression of innexins in the yellow fever mosquito, Aedes aegypti: insights into putative life stage- and tissue-specific functions of gap junctions

Published in: Calkins, T.L., Woods-Acevedo, M.A., Hildebrandt, O., & Piermarini, P.M. 2015. The molecular and immunochemical expression of innexins in the yellow fever mosquito, Aedes aegypti: Insights into putative life stage- and tissue-specific functions of gap junctions. Comparative Biochemistry and Physiology Part B: Biochemistry and Molecular Biology, 183, 11–21. doi:10.1016/j.cbpb.2014.11.013

Abstract

Gap junctions (GJ) mediate direct intercellular communication by forming channels through which certain small molecules and/or ions can pass. Connexins, the proteins that form vertebrate GJ, are well studied and known to contribute to neuronal, muscular and epithelial physiology. Innexins, the GJ proteins of insects, have only recently received much investigative attention and many of their physiological roles remain to be determined. Here we characterize the molecular expression of six innexin

(Inx) genes in the yellow fever mosquito Aedes aegypti (AeInx1, AeInx2, AeInx3, AeInx4,

AeInx7, and AeInx8) and the immunochemical expression of one innexin protein, AeInx3, in the alimentary canal. We detected the expression of no less than four innexin genes in each mosquito life stage (larva, pupa, adult) and tissue/body region from adult males

51 and females (midgut, Malpighian tubules, hindgut, head, carcass, gonads), suggesting a remarkable potential molecular diversity of GJ in mosquitoes. Moreover, the expression patterns of some innexins were life stage and/or tissue specific, suggestive of potential functional specializations. Cloning of the four full-length cDNAs expressed in the

Malpighian tubules of adult females (AeInx1, AeInx2, AeInx3, and AeInx7) revealed evidence for 1) alternative splicing of AeInx1 and AeInx3 transcripts, and 2) putative N- glycosylation of AeInx3 and AeInx7. Finally, immunohistochemistry of AeInx3 in the alimentary canal of larval and adult female mosquitoes confirmed localization of this innexin to the intercellular regions of Malpighian tubule and hindgut epithelial cells, suggesting that it is an important component of GJ in these tissues.

1. Introduction

Gap junctions (GJ) are intercellular channels that allow for direct communication between adjacent cells via the transport of small molecules and ions (Cao et al. 1998).

GJ are formed when a hemichannel from one cell docks with a hemichannel in an adjacent cell (Phelan and Starich 2001; Johnson et al. 2012; Scemes et al. 2009). In arthropods, the hemichannels of GJ are hexamers composed of ‘innexin’ protein subunits, whereas in mammals and other vertebrates the hemichannels are hexamers composed of ‘connexin’ protein subunits (P Phelan et al. 1998; Swenson et al. 1989).

Although innexin and connexin genes are evolutionarily distinct and encode proteins

52 that share nominal amino acid homology, their encoded proteins have convergently evolved many structural and functional features (Abascal and Zardoya 2013).

In vertebrates, GJ play important roles in embryogenesis, neurophysiology, muscle physiology, epithelial physiology, and endocrinology (Bosco et al. 2011). In arthropods, the physiological roles of GJ have not yet been fully elucidated, but they appear to play important roles in development, reproduction, and epithelial physiology.

In Drosophila melanogaster, innexin 1 (ogre) is critical to the development of retinal neurons (Curtin et al. 2002), and innexin 3 is necessary for dorsal closure in embryogenesis (Giuliani et al. 2013). Moreover, innexin 4 (zero population growth or zpg) is involved with functional gonad formation in males of both D. melanogaster flies and Anopheles gambiae mosquitoes (Magnusson et al. 2011; Tazuke et al. 2002), and is necessary for early germ cell differentiation in female D. melanogaster (Gilboa et al.

2003). Furthermore, innexins are likely responsible for the functional coupling of epithelial cells in the renal (Malpighian) tubules of mosquitoes (K W Beyenbach and

Piermarini 2011; Piermarini and Calkins 2014; Weng et al. 2008).

The goal of this study was to describe the post-embryonic expression of the six putative innexin (AeInx) genes in the yellow fever mosquito A. aegypti (Weng et al.

2008) and to localize the expression of one innexin protein (AeInx3) in the alimentary canal. We hypothesized that innexins would exhibit different patterns of mRNA expression throughout the mosquito life cycle and among various tissues, providing insights into the putative functional roles of these genes in mosquitoes. Furthermore,

53 we hypothesized that the immunoreactivity of AeInx3 would concentrate near the intercellular regions of epithelial cells of the alimentary canal where it would potentially form GJ.

2. Materials and Methods

2.1 Mosquitoes

Eggs of Aedes aegypti were obtained through the Malaria Research and

Reference Reagent Resource Center (MR4) as part of the BEI Resources Repository

(Liverpool strain; LVP-IB12 F19, deposited by M.Q. Benedict). Mosquitoes were reared as described in Piermarini et al. (2011) in an environmental chamber set at 28C and

80% relative humidity with a 12 h:12 h light:dark cycle.

2.2 Dissections:

Adult and larval mosquitoes were anesthetized on ice, and then grasped with forceps (Dumont #5; Fine Science Tools, Inc., Foster City, CA) either at the thorax to remove the legs and heads (adults) or one segment posterior to the head (larvae) to remove the head. The insect was then submerged in a mosquito Ringer solution consisting of the following in mM: 150 NaCl, 3.4 KCl, 1.7 CaCl2, 1.8 NaHCO3, 1 MgCl2, 5

Glucose, 25 HEPES. The pH of the Ringer solution was adjusted to 7.1 with NaOH before use and the desired osmolality was verified (330 ± 5 mOsm/kg) with a vapor pressure

54 osmometer (Wescor, Logan, UT). The second to last abdominal segment was grasped with forceps and gently pulled away from the rest of the abdomen, bringing with it the alimentary canal and gonads. Tissues of interest were isolated and placed into separate

1.5 ml sterile microcentrifuge tubes (USA Scientific; Orlando, FL) containing the mosquito Ringer solution on ice. For whole insect samples, dissection was unnecessary.

Once all mosquito samples were collected, the Ringer solution was aspirated and replaced with TRIzol (Life Technologies, Carlsbad, CA) before storing at -80C.

2.3 Qualitative RT-PCR

Using an approach modified from Piermarini et al. (2013) and Rouhier and

Piermarini (2014), qualitative RT-PCR was performed to assess Aedes aegypti innexin

(AeInx) gene expression. In brief, total RNA was isolated from preserved tissues or whole insects using the method of Chomczynski and Sacchi (1987). The cDNA templates were synthesized using 1 g of total RNA and GoScript Reverse Transcriptase (Promega,

Madison, WI) with random hexamers, following the manufacturer’s protocol. Primers for the RT-PCR reactions were designed against the six predicted innexin genes in the A. aegypti genome (i.e., AeInx, Table 4) using the SciTools-Real-Time PCR application on the Integrated DNA Technologies web site (www.idtdna.com). Note that since the original description of these genes in A. aegypti (Weng et al. 2008), one has been renamed. That is, the gene initially designated as ‘passover’ due to its homology to the shakB/passover gene in D. melanogaster is now named as innexin 8 (Inx8) to follow the

55 nomenclature of Hasegawa and Turnbull (2014). The AeInx primers were designed to amplify a product of approximately 500 base pairs (bp). A 300 bp amplicon of the ribosomal protein S7 gene (RPS7) was used as the internal control (Piermarini et al.

2013). All PCR primers flanked at least one intron to potentially detect the amplification of genomic DNA.

Table 4: RT-PCR primers used to assess the expression of innexin genes in A. aegypti. Accession numbers are from Vectorbase (www.vectorbase.org).

Innexin Accession Forward primer Reverse primer

number AeInx1 AAEL014846 CCGTATTCCGCCTCCACAATAG ACCAAAGTGCTTCACATGC

AeInx2 AAEL014847 AAGATCACGCCTGTCAGAAC GGTTGAACTTAGGGATTGGA

AeInx3 AAEL011248 CAGCTACTCGTTCGGTTACTTC ATGTCCTCACGTTGATGTTCTGTCCC

AeInx4 AAEL006726 TTCTGTTGGACACTGGGAAC CCAGAGAATTATACGAAGT

AeInx7 AAEL008588 CCAGGCAATACATAGGAGAGC CCAGGTCGCAGAGTACGTGTTTC

AeInx8 AAEL014227 GAAGTACCCTATCCAGGTGTT CCGCTGACACCATACTTGTA

G G

All RT-PCR reactions were performed using GoTaq Green 2x MasterMix

(Promega, Madison, WI). Each reaction consisted of the following: 25 L GoTaq Green, 1

L AeInx primer pair (10 M), 1 L RPS7 primer pair (10 M), 1 L cDNA, 2 or 4 L of 25 mM MgCl2 and nuclease free water up to 50 l. Reactions were individually optimized for MgCl2 concentration; reactions for AeInx1, AeInx3, and AeInx7 used 4 mM MgCl2, whereas those for AeInx2, AeInx4, and AeInx8 used 5 mM MgCl2.

The amplification protocol consisted of a denaturation at 95 C for 5 min, followed by 50 cycles of 95C for 30 s, 52C or 58C for 15 s, and 72C for 30 s. Each reaction was performed in triplicate. AeInx1 and AeInx8 reactions used an annealing temperature of 52C, whereas AeInx2, AeInx3, AeInx4 and AeInx7 used an annealing temperature of 58C.

PCR products were separated by electrophoresis on a 1% agarose gel for 30 minutes at 100 mV, and visualized via ethidium bromide staining on a Gel Logic Imager

(Kodak, Rochester, NY). The identities of all PCR products were confirmed by sequencing at the Molecular and Cellular Imaging Center at the Ohio Agricultural Research and

Development Center (OARDC) of The Ohio State University, Wooster, OH.

In order to qualitatively assess the expression of the AeInx genes, we measured the optical densities of the AeInx bands and compared them to RPS7. For a given gel, the optical density of the AeInx and RPS7 bands were quantified using ImageJ pixel counting software (NIH). For each AeInx band in a given tissue, the mean optical density for 3 replicates was compared to that of RPS7 using a student’s T-test. If the mean optical density of an AeInx band was significantly lower than the mean RPS7 value, then it was categorized as ‘weak’ expression. If mean optical density of an AeInx band was not significantly different than the mean RPS7 value, then it was categorized as

‘moderate’ expression. If mean optical density of an AeInx band was significantly greater than mean RPS7 value, then it was categorized as ‘strong’ expression. If no detectable

57 band was present in all 3 replicates, then it was categorized as ‘not detectable’. The categorizations are graphically summarized in Figures 12-14.

2.4 Cloning

Malpighian tubules were dissected from adult female mosquitoes in Ringer solution and RNA was isolated from Malpighian tubules, as described above. The RNA was used as a template to generate 5’ and 3’ single-stranded cDNA libraries with the

GeneRacer Kit (Invitrogen, Carlsbad, CA), as described previously (Piermarini et al.

2010). The resulting cDNA libraries were used for 5’ and 3’ rapid amplification of cDNA ends (RACE), respectively, as described previously (Piermarini et al. 2010).

RACE was performed for each innexin using one gene-specific primer (Table 5) and a generic ‘Generacer’ primer, according to the manufacturer’s instructions. In brief, each RACE reaction consisted of 0.5 L cDNA, 1.5 L RACE primer, 0.5 l gene specific primer, and 22.5 L Platinum PCR High Fidelity SuperMix (Invitrogen), and was subjected to a touchdown RT-PCR amplification, following the manufacturer’s protocol.

Products from RACE reactions were ligated into a TOPO 4.1 plasmid (Invitrogen) and transformed into Premade Z-Competent E. coli Cells (Zymo Research, Irvine, CA).

The E. coli cells were spread on a 2% agar LB media plate. After overnight growth, individual colonies were picked and grown up in liquid LB media at 37C in a shaking incubator overnight. The plasmid DNA was extracted using a QIAGEN QIAprep Spin

Miniprep kit (Qiagen, Valencia, CA). Plasmid DNA was sequenced at the Molecular and 58

Cellular Imaging Center at the Ohio Agricultural Research and Development Center of

The Ohio State University, Wooster, OH. Full-length cDNA sequences were assembled from the RACE sequencing data using CLC Workbench 6 Software. The consensus sequences were built from five 5’ and seven 3’ RACE sequences for AeInx1, four 5’ and nine 3’ RACE sequences for AeInx2, six 5’ and six 3’ RACE sequences for AeInx3, and five

5’ and three 3’ RACE sequences for AeInx7. Consensus sequences were deposited into

Genbank and assigned accession numbers as shown in Table 6.

Table 5: Gene specific primers used for RACE.

Innexin Forward primers (3’-RACE) Reverse primers (5’-RACE)

1F: 1R: GTATTAACGAGAGGTAGGACGGGAA GGCGAGTTCCGCTACAACATCCTTGA AeInx1 2F: 2R: TTTATGGGATGCAACGGAAGGAGGA GGGAATACGTAAACCATCGGATCTAA 1F: 1R: AGTCACGTCGACGGCCATGATGAAG AAACGGTAGACCAATGATGCACCAG AeInx2 2F: 2R: CATCTGCTGTGCATAGAACCTAGAAA TCTTTGCCCTCAAATTTGAGAGAGA 1F: 1R: CGTGTTCAGATTCGACGGACAAGACT AGTGAATCGTAGAGGTACTTAGCGAC 2F: 2R: AeInx3 CTTCGGTAGCACACTCCGGTCTCGCC CAGGATGAACCAGAACCACAGGAAG 3F: C GGATCGCAACTCCGTAGAGGCACCC 1F: 1R: CGTCTTGTGGATGGTCTTCGTATGGT ATTCCAGGACTAACCTTGCAGAAAA 2F: 2R: AeInx7 CCTCACCAGGAGTTGGACTAACATA ATAAGTTCAGTTTCGTCAGCCTCAT 3R: CCAGTCCGAAAATGAAAGGCTCTCCG

Table 6: Genbank accession numbers for cloned innexin cDNAs and their splice variants.

Innexin cDNA Accession number Inx1-A KJ736822 Inx1-B KJ736823 Inx2 KJ736824 Inx3-A KJ736825 Inx3-B KJ736826 Inx7 KJ736827

2.5 Sequence Alignment

Deduced amino-acid sequences of the cloned innexins were aligned using

ClustalW2 (European Bioinformatics Institute, Hinxton, Cambrigeshire, UK).

Transmembrane domains were predicted using a Kyte-Doolittle hydrophobicity analyses

(window size 19; Figure 11) with supporting analysis from 1) Phobius prediction of transmembrane topology and signal peptide software (European Bioinformatics

Institute) and 2) previous predicted membrane topologies for innexins of Drosophila melanogaster and Bombyx morii (S. M. Hong et al. 2009).

Figure 11: Kyte-Doolittle mean hydrophobicity plot for innexins 1, 2, 3 and 7. Window size 19. Innexins 1, 2, 3 and 7 are represented by the red, blue, green and brown lines, respectively.

2.6 Anti-Inx3 Antibody

To raise and affinity purify a rabbit polyclonal antibody against an AeInx3 synthetic peptide, we hired 21st Century Biochemicals (Marlboro, MA). The peptide for antibody production and affinity purification corresponded to the COOH-terminal domain of the predicted AeInx3-A protein (L374EMAPIYPEIGKYGKDTAI; Figure 5).

Notably, the first 14 residues of this peptide are perfectly conserved between AeInx3-A and AeInx3-B, but the last six residues are only conserved AeInx3-A. Thus, the resulting antibody may have a higher affinity for the AeInx3-A vs. the AeInx3-B isoform.

2.7 Western Blotting

Whole animals for Western blot analysis were lysed in 10% Ringer solution.

Protein concentration was measured using the Pierce BCA protein assay (Thermo

Scientific, Rockford, IL) and then an equal volume of urea lysis buffer was added to the sample as well as an appropriate volume of a 5x Laemmli sample buffer (Laemmli 1970;

Piermarini et al. 2010). Proteins were separated via denaturing sodium dodecyl sulfate

(SDS) polyacrylamide gel electrophoresis (PAGE) through a 6% stacking/12% resolving gel at 125 mV for 2.5 hours in an XCell SureLock mini cell electrophoresis box

(Invitrogen). Proteins were transferred to a polyvinylidene difluoride (PVDF) membrane

(BioRad) in the XCell II Blot module (Invitrogen) at 35 mV for 2 hours. Transfer of protein was confirmed by staining the PVDF membrane with Ponceau S. Membranes were then blocked for 2 hours in 5% nonfat dry milk (Nash Finch Co., Minneapolis MN) in Tris

Buffered Saline with 0.1% Tween 20 (TBST) at room temperature on a rocking incubator.

Membranes were incubated overnight with the affinity-purified anti-AeInx3 antibody

(0.4  C. The membrane was washed 3 times in

TBST for 5 minutes each. The membrane was then incubated in secondary antibody, horseradish peroxidase-conjugated goat anti rabbit IgG (Thermo Scientific) diluted

1:20,000 at room temperature for 2 hours. After an additional 3 washes in TBST the membrane was incubated in SuperSignal West Dura (Thermo Scientific) for 5 minutes, transferred to clean cellophane wrap and imaged using a My ECL imager (Model

62236X, Thermo scientific).

2.8 Whole Mount Immunolabeling

Tissues for whole mount immunohistochemistry were dissected from adult or larval mosquitoes as described in section 2.2. After dissection, tissues of interest were transferred to a clean 1.5 ml centrifuge tube and fixed for 45 minutes in 1 ml of a fixative solution containing 4% paraformaldehyde (Electron Microscopy Sciences,

Hatfield, PA) in phosphate-buffered saline (PBS). Tissues were washed for 5 minutes with 1 ml of PBS three times, followed by two 10-minute washes of 0.1% TritonX 100

(Fisher Scientific, Pittsburg, PA) in PBS and three additional 5-minute washes with 0.1%

Tween 20 (Fisher Scientific) in PBS (PBST). Tissues were blocked for 1.5 hours at room temperature with 10% normal goat serum (Thermo Scientific, Rockford, IL) supplemented with casein (Vector Laboratories, Burlingame, CA) in PBST. Tissues were incubated with the rabbit anti-AeInx3 primary antibody (2.3 µg/ml) diluted in PBST supplemented with casein overnight at 4° C.

On the following day, tissues were washed three times with PBST (5 min each) and then incubated in a 1:600 dilution of Dylight 488 IgG goat anti-rabbit secondary antibody (Thermo Scientific) in PBST supplemented with casein for 1.5 hours at room temperature. Tissues were finally incubated with DAPI (Thermo Scientific, Rockford, IL) at a 1:5000 dilution. Peptide blocked tissue preparations followed the same procedure with the exception that the primary antibody was pre-incubated for 1 hour at 37 C with the immunogenic peptide in 10-fold excess. Tissues were imaged using a Leica TCS SPS II confocal microscopy system along with the Leica Microsystems LAS AF software (Leica

Microsystems, Buffalo Grove, IL) at the Molecular and Cellular Imaging Center of the

Ohio Agricultural Research and Development Center (Wooster, OH).

3. Results

3.1 Patterns of AeInx expression:

To describe the expression of the six predicted AeInx genes across developmental stages we conducted RT-PCR using gene-specific primers for each AeInx and an internal positive control gene (RPS7). As explained in the Materials and Methods section, we qualitatively categorized the expression of each innexin relative to RPS7 as not detectable, weak, moderate, or strong.

Figure 12 shows that larvae (4th instar) are characterized by moderate or strong expression of AeInx1, AeInx2, AeInx3 and AeInx7, and weak expression of AeInx8

(expression of AeInx4 is not detectable). Pupae exhibit a similar pattern of AeInx expression as larvae, but are distinguished from larvae by strong expression of AeInx2 and moderate expression of AeInx8 (Figure 12). Adult males are characterized by strong expression of AeInx2 and moderate expression of AeInx1, AeInx3 and AeInx7 (expression of AeInx4 and AeInx8 are not detectable). On the other hand, adult females are characterized by the strong expression of AeInx1, AeInx2, AeInx3, and AeInx7, and moderate expression of AeInx4 (expression of AeInx8 is not detectable).

Figure 12: Histogram summarizing the qualitative expression of each innexin in mosquito life stages and sexes. Numbers indicate the corresponding innexin gene (e.g., 1 = AeInx1). The qualitative score for each innexin was assigned based on the mean optical density of an AeInx PCR product relative to that of internal control gene RPS7, as described in the Materials and Methods

Figure 13 compares the expression of AeInx genes among three epithelial tissues of the alimentary canal (Malpighian tubules, midgut, and hindgut) in adult female and male mosquitoes. Common to all three tissues examined for both sexes is the expression of AeInx2, AeInx3, and AeInx7 at moderate or strong levels; one exception is the adult female midgut, which expresses AeInx3 weakly. The expression of AeInx1 is

65 weak in all three tissues of females, but is moderate in those of males. The expression of

AeInx4 is not detectable in all three tissues of females and the midgut of males, but is present at weak and moderate levels in the Malpighian tubules and hindgut, respectively, of males. Moreover, the expression of AeInx8 is not detectable in the

Malpighian tubules and midgut of females and males, but is weak in the hindgut of males and females.

Figure 13: Histogram summarizing the qualitative expression of each innexin in tissues of the alimentary canal of adult female and male mosquitoes. Numbers indicate the corresponding innexin gene (e.g., 1 = AeInx1). The qualitative score for each innexin was assigned based on the mean optical density of an AeInx PCR product relative to that of internal control gene RPS7, as described in the Materials and Methods.

Figure 14 compares the expression of AeInx genes among the head, carcass, and gonads (ovaries or testes). Common to all of these tissues in both sexes is the moderate

66 or strong expression of AeInx1, AeInx2, and AeInx3. Notably, the expression of AeInx4 is not detectable in the head of females, but is moderate in those of males, and expressed at moderate or strong levels in the other tissues with the exception of the male carcass where it is weak. The expression of AeInx7 is weak and moderate in the head of females and males, respectively, while it is strong in the carcass of both females and males.

AeInx7 expression is not detectable in gonads of females (ovaries), and is weakly expressed in those of males (testes). AeInx8 is expressed at weak (females) or moderate

(males) levels in all three tissues, with the exception of the testes where its expression is not detectable.

Figure 14: Histogram summarizing the qualitative expression of each innexin in the head, carcass, and gonads. Numbers indicate the corresponding innexin gene (e.g., 1 = AeInx1). The qualitative score for each innexin was assigned based on the mean optical density of an AeInx PCR product relative to that of internal control gene RPS7, as described in the Materials and Methods.

3.2 AeInx cDNAs and Gene Structures

Our laboratory is especially interested in the putative roles of innexins in the synchronization and regulation of renal functions in adult female mosquitoes

(Piermarini and Calkins 2013). Thus, we aimed to clone the AeInx cDNAs (and any splice variants) expressed in female Malpighian tubules (AeInx1, AeInx2, AeInx3, and AeInx7;

Figure 15) and elucidate their respective gene structures.

Figure 15A shows the structure of the AeInx1 gene, based on cloned cDNAs.

AeInx1 consists of six exons spanning ~34 kb on ‘supercontig 1.1293’ of the A. aegypti genome, and is expressed as two distinct splice variants in the Malpighian tubules:

AeInx1-A and AeInx1-B. The AeInx1-A cDNA consists of 1,510 bp whereas the AeInx1-B cDNA consists of 1,040 bp. The variants differ in their 3’ ends where AeInx1-A has a sixth exon not found in AeInx1-B. Notably, in AeInx1-A, the fifth exon is slightly truncated, resulting in a loss of the stop codon that occurs in AeInx1-B, which extends the open reading frame of AeInx1-A into exon 6 and lengthens its 3’ UTR, compared to AeInx1-B.

The predicted ORF of AeInx1-A consists of 1,092 bp encoding 364 amino acids. In contrast, the predicted ORF of AeInx1-B consists of 867 bp encoding 289 amino acids.

Both predicted proteins are identical with the exception of the COOH terminal domain, which is 75 amino acids longer in AeInx1-A (Figure 16).

Figure 15: Mapping of the AeInx1, AeInx2, AeInx3, and AeInx7 genes (A, B, C, and D respectively). The top drawing of each panel shows the gene structure based on the cloned cDNAs with the thick, horizontal orange bars representing exons and the thin black lines representing introns. The bottom drawings of each panel show cartoons of the exon composition(s) for cloned cDNAs from each gene. Cyan-shaded areas represent the open reading frames and red areas represent the untranslated regions. Splice variants are indicated as ‘A’ or ‘B’.

Figure 16: Alignments of the deduced amino-acid sequences encoded by the AeInx1 and AeInx3 mRNA splice variants. Differential splicing in Inx1 results in identical proteins with the exception of a longer COOH-terminal domain in AeInx1A. Differential splicing in Inx3 results in identical proteins for the first 386 amino acids, but differences in the identities of 5 of the last 6 mutually amino acids of AeInx3-A and a longer COOH- terminal domain in AeInx3-B indicated in grey.

Figure 15B shows the structure of the AeInx2 gene, based on the cloned cDNA.

AeInx2 consists of only two exons, which span ~7 kb of ‘supercontig 1.293’, ~20 kb

70 downstream from AeInx1; we did not detect the expression of AeInx2 splice variants.

The AeInx2 cDNA consists of 2,586 bp with a predicted ORF of 1,077 bp encoding 359 amino acids. The predicted 5’-UTR, ORF, and a partial 3’-UTR are all contained within exon 1, while exon 2 contains the remainder of the 3’-UTR.

Figure 15C shows the structure of the AeInx3 gene, based on the cloned cDNAs.

AeInx3 consists of six exons spanning ~19kb of ‘supercontig 1.560’, and we detected the expression of two splice variants: AeInx3-A and AeInx3-B. The first exon of AeInx3 resides ~15 kb upstream from the subsequent five exons (Figure 15C); the latter 5 exons reside within a 3 kb range. The AeInx3-A cDNA consists of 2,774 bp with a predicted ORF of 1,176 bp encoding 392 amino acids. The AeInx3-B cDNA is ~500 bp longer, consisting of 3,209 bp with a predicted ORF of 1,197 bp encoding 399 amino acids. While there is a

500 bp difference in the cDNAs, both predicted proteins are identical with the exception of the extreme COOH terminal domains (Figure 16). This is due to the retention of the intron between exons 5 and 6 in the ‘A’ variant and an extension of the sixth exon in the

‘B’ variant.

Figure 15D shows the structure of the AeInx7 gene, based on the cloned cDNA.

AeInx7 consists of four exons spanning ~11 kb of ‘supercontig 1.333’, and we did not detect the expression of splice variants. The AeInx7 cDNA consists of 1,350 bp and a predicted ORF of 1,221 bp encoding 407 amino acids, with relatively short 5’ and 3’

UTRs.

3.3 Predicted Innexin Proteins:

Figure 17 shows an amino-acid sequence alignment of the deduced AeInx1-A,

AeInx2, AeInx3-A, and AeInx7 proteins, based on the cloned cDNAs. All four innexins exhibit the expected, predicted membrane topology with relatively short and long cytosolic NH2-terminal and COOH-terminal domains (NT and CT, respectively, in Figure

17) that flank a transmembrane domain containing 4 membrane-spanning segments

(TM1-TM4 in Figure 6). Also notable are 1) the perfectly conserved innexin signature

‘YYQWV’ motif, located at the start of TM2 in all four of the innexins, and 2) the highly conserved extracellular loop (EL2 in Figure 17) between TM3 and TM4, which includes two cysteine residues (‘*’ in Figure 17) that are highly conserved across connexins and innexins (Barnes 1994; P Phelan and Starich 2001). These cysteine residues play important roles in the docking of connexin hemichannels between the plasma membranes of neighboring cells (X. Bao et al. 2004). Overall, the proteins encoded by the different mosquito innexin cDNAs share only 30-50% amino-acid identity with each other, and share more identity with their respective orthologs in Drosophila melanogaster (Table 7).

Figure 17: Amino-acid sequence alignment of the four innexins cloned from adult female Malpighian tubules. Black shaded regions indicate identical residues among the innexins, whereas gray shaded regions indicate similar residues among the innexins. The transmembrane regions as predicted by Kyte-Doolittle hydropathy plots (Figure 11) in association with predicted membrane topologies of D. melanogaster and B. mori innexins (S.-M. Hong et al. 2008). The key features of the proteins are labeled, NH2- terminus (NT), transmembrane region (TM), conserved cysteine residues (*) extracellular loop (EL), intracellular loop (IL) and the COOH-terminus (CT).

Table 7: Percent amino-acid identity of the cloned innexins and splice variants to each other and their closest ortholog in D. melanogaster.

Inx1-A Inx1-B Inx2 Inx3-A In3-B Inx7 Inx1-A 100 45.1 41.8 41.8 33.2 Inx1-B 100 47.1 46.4 46.4 38.1 Inx2 45.1 47.1 46.0 46.0 39.3 Inx3-A 41.8 46.4 46.0 98.5 32.4 In3-B 41.8 46.4 46.0 98.5 31.6 Inx7 33.2 38.1 39.3 32.4 31.6 D. melanogaster 72.4 78.2 81.8 68.0 67.5 48.3

A PROSCAN analysis of the AeInx amino-acid sequences identified a predicted N- glycosylation site in the second extracellular loop of both AeInx3 variants (N229RSD), and the first extracellular loop of AeInx7 (N84ESM). The PROSCAN analysis (npsa-pbil.ibcp.fr) also revealed putative phosphorylation sites (Table 8). Aeinx1-A has one predicted phosphorylation site in the COOH-terminus, which is not present in AeInx1-B, due to the truncated COOH-terminus of AeInx1-B. AeInx2 has five putative phosphorylation sites, one of which is located in the NH2-terminus; the other four reside in the C-terminus.

AeInx3-A and AeInx3-B share the same five putative phosphorylation sites; one is in the intracellular loop and four are in the COOH-terminus. AeInx7 has the most putative phosphorylation sites with nine. Two reside in the intracellular loop and the remaining seven are in the COOH-terminus.

Table 8: Phosphorylation sites as identified by PROSCAN analysis.

Residue Location Residue Identity AeInx1A 297-299 SFR AeInx1B N/A N/A AeInx2 7-9 SVK 310-313 SSK 319-321 SLK 335-342 KNIDPLIY 350-352 SEHD AeInx3A/B 157-160 SPDE 303-305 TTR 315-318 TAPD 347-350 SYSE 372-375 STLE AeInx7 176-179 SELD 150-157 RMVEVSRY 319-321 SVK 348-351 SFSD 381-384 TRHE 385-387 SSK 386-388 SKK 387-390 KKIS 404-406 SEK

3.4 Immunochemical expression of AeInx3

Using an affinity-purified antibody, we characterized the immunochemical

expression of AeInx3 in the alimentary canal of adult females and 4th instar larvae. The

predicted molecular mass of AeInx3 is ~45 kDa. In crude lysates of adult female

mosquitoes, the anti-AeInx3 antibody detected several bands, from as low as ~49 kDa to

as high as >130 kDa (Figure 18). The multiple bands suggest the presence of

glycosylated and/or oligomeric forms of AeInx3.

Figure 18: Western blot of AeInx3 immunoreactivity in a crude lysate of adult female mosquitoes. Expected size of the predicted AeInx3 protein is ~45 kDa.

Figure 19 shows representative localizations of AeInx3 immunoreactivity in the alimentary canal of adult females. In the proximal end of the Malpighian tubules, where there are no stellate cells, AeInx3 immunoreactivity is prominent at the intercellular boundaries between principal cells (Figure 19A, arrow). Diffuse, punctate labeling in the cytoplasm is also present. In the distal end of the tubules, where stellate cells intercalate among principal cells, the AeInx3 immunoreactivity occurs in principal cells at the intercellular boundaries between principal cells (not shown), and as dense, punctate labeling in the cytoplasm of principal cells (Figure 19B). AeInx3 immunoreactivity is not detectable in the cytoplasm of stellate cells (Figure 19B, arrowhead), but is found in principal cells that neighbor stellate cells (Figure 19B, arrow).

Figure 19: Localization of AeInx3-immunoreactivity (green) in isolated tissues of the adult female alimentary canal. (Continued on next page)

Figure 19 continued: Nuclei are counterstained with DAPI (cyan). A) Proximal end of an isolated Malpighian tubule. The white arrow indicates AeInx3 localization to the intercellular membranes between principal cells. B) Distal, blind end of an isolated Malpighian tubule. Arrowhead indicates a stellate cell lacking AeInx3 signal and arrow indicates AeInx3 immunoreactivity in a neighboring principal cell. C) A midgut showing no detectable AeInx3 localization. D) A midgut showing immunoreactivity in associated tracheal tubes (arrow). E) Posterior hindgut showing the rectum (arrow) and distal end of ileum (arrowhead). F) Anterior hindgut showing the ileum; inset shows digital zoom of the region outlined by the white square.

In the midgut of adult females, no immunoreactivity is detectable in the epithelium (Figure 19C), but the tracheal tubes associated with the midgut exhibit detectable labeling (Figure 19D, arrow). In the hindgut, strong labeling of the rectum

(Figure 19E, arrow) and ileum (Figure 19E, arrowhead; Figure 19F) occurs at the intercellular boundaries and within the cytoplasm of the epithelial cells. Figure 20E shows the junction of the midgut and hindgut where the Malpighian tubules attach, which demonstrates that the relative overall immunoreactivities exhibited by these three tissues in adult females are consistent with the respective, relative mRNA expression patterns observed in Figure 13. Similar patterns of AeInx3 immunoreactivity and localization are observed in the respective isolated tissues from larvae (Figure 20 A-

C). Preabsorbing the anti-AeInx3 antibody with its antigenic peptide depleted the observed immunolabeling (e.g., Figure 20F). In adult male tissues, AeInx3 immunoreactivity was similar to those of females (Data not shown).

Figure 20: AeInx3 localization (green) in larval (A-C) and adult female (D-F) tissues. Nuclei are counterstained with DAPI (cyan) in panels A-C and E. A) midgut; the arrow indicates localization associated with the tracheal tubes. B) Malpighian tubule; arrowheads indicate stellate cells without immunoreactivity, and arrows indicate intercellular regions of neighboring principal cells. C) Hindgut; inset shows digital zoom of the region outlined by the white square. (Continued on next page) 79

Figure 20 continued: D) Alimentary canal; open arrow, filled arrow, and arrowhead indicate the anterior hindgut, Malpighian tubules, and midgut, respectively. E) Malpighian tubule incubated with anti-AeInx3 antibody after pre-adsorption with the immunogenic AeInx3 peptide.

4. Discussion

This study is the first to compare and characterize the expression of innexins throughout the mosquito life cycle and in isolated tissues of mosquitoes. As discussed below, our results reveal trends in innexin mRNA expression that shed light on the potential developmental and physiological roles innexins play in mosquitoes. However, it is important to emphasize that these trends do not reveal the actual physiological function of individual innexins, but rather provide us with a starting point for developing hypotheses on their putative roles.

4.1 General trends

AeInx1, AeInx2, AeInx3, and AeInx7 mRNAs are expressed consistently across all life stages and sexes examined (Figure 12), which suggests that these innexins are the most abundantly expressed in A. aegypti and may contribute to essential house-keeping functions throughout the mosquito life cycle. In contrast, AeInx4 and AeInx8 are the least consistently expressed, suggesting potential life stage- and/or sex-specific roles.

Studies examining innexin expression in Bombyx mori have demonstrated that BmInx2,

BmInx3 and BmInx4 exhibit similar expression patterns throughout the life cycle as 80 those reported here in the mosquito (S.-M. Hong et al. 2008; S. M. Hong et al. 2009).

Thus, the developmental expression patterns of some innexins may be well conserved across holometabolous insects.

4.2 A potential role of AeInx4 in reproduction

Adult females are the only life stage and sex in which the expression of AeInx4 mRNA is detectable in the whole animal (Figure 12), suggesting a sex-specific function of this innexin. Our analysis of isolated tissues shows that this gene is expressed primarily in the carcass and ovaries of adult females (Figure 3). The carcass consists of the thoracic and abdominal body walls, which contain the flight musculature, its innervation, and the fat body. Given that the fat body plays a key role in vitellogenesis of female mosquitoes after a blood meal, the expression of AeInx4 in the carcass may indicate a role of this innexin in fat body-mediated vitellogenesis. Consistent with this notion, the expression of AeInx4 is weak in the carcass of males, which do not feed on blood or undergo vitellogenesis.

Our finding of AeInx4 mRNA expression in the ovaries of females is consistent with previous studies, which have detected expression of Inx4 in the ovaries of D. melanogaster and B. mori (Bohrmann and Zimmermann 2008; S.-M. Hong et al. 2008; S.

M. Hong et al. 2009). Furthermore, the coexpression of AeInx4 with AeInx1, AeInx2, and

AeInx3 in A. aegypti ovaries (Figure 14) mirrors that of D. melanogaster ovaries, which also express these innexins (Bohrmann and Zimmermann 2008). Notably, in the ovaries

81 of D. melanogaster, Inx4 immunoreactivity colocalizes with that of Inx2 where it may form connections between soma and germ-cell lines (Bohrmann and Zimmermann

2008). Moreover, in the ovaries of D. melanogaster, Inx4 and Inx2 are necessary for 1) germ cell differentiation and survival, and 2) cyst and egg chamber formation (Gilboa et al. 2003; Mukai et al. 2011). Interestingly, AeInx4 mRNA expression is also strong in the testes of males (Figure 14), but it must not be abundant enough for its detection in the whole animal (Figure 12). The finding of strong expression in the testes is consistent with previous functional genetic studies in A. gambiae and D. melanogaster, which have shown that the disruption of Inx4 expression elicits a sterile male phenotype

(Magnusson et al. 2011; Tazuke et al. 2002). Thus, AeInx4 may play similar key roles in the reproductive biology of A. aegypti.

4.3 A potential role of AeInx8 in development and neuromuscular function:

AeInx8 mRNA expression is detectable at weak and moderate levels, in larval and pupal life stages, respectively, but it is not expressed at detectable levels in adults

(Figure 12). This suggests that AeInx8 has a post-embryological role in larval development and pupal metamorphosis. In pupal D. melanogaster, Inx8 expression occurs in neuronal tissue and decreases within two days after eclosion into adults

(Crompton et al. 1995). Thus, a similar phenomenon may occur in mosquitoes.

Interestingly, in adult mosquitoes, AeInx8 mRNA expression is detectable consistently at weak or moderate levels in the hindgut, head, and carcass of both males

82 and females, and in the ovaries of females (Figures 13-14), which suggests that AeInx8 expression persists throughout the adult stages, but at levels that are not detectable in the whole animal. The expression of AeInx8 in the head and carcass is expected as both contain neuronal tissues (e.g., ganglia, nerve cord), and Inx8 orthologs of other insects

(D. melanogaster, Schistocerca gregaria) have been implicated in the connectivity of the giant fiber system and frontal ganglion (Anava et al. 2009; Krishnan et al. 1993).

The finding of AeInx8 mRNA expression in the hindgut, but not the midgut and

Malpighian tubules (Figure 13) is intriguing, because these are all epithelial tissues involved with transepithelial ion and water transport. However, the hindgut is exceptional in that it undergoes spontaneous, peristaltic muscle contractions, even when isolated from the whole animal (Kwon and Pietrantonio 2013; Messer and Brown

1995). Thus, it is tempting to speculate that AeInx8 may be playing a role in coordinating these peristaltic movements. Consistent with this notion, AeInx8 mRNA is expressed in the ovaries (and not the testes) and the associated oviduct is known to undergo similar spontaneous contractions as the hindgut (Kwon and Pietrantonio 2013; Messer and

Brown 1995). Thus, AeInx8 may play an important role in the excretory and reproductive physiology of mosquitoes by modulating muscle function in the hindgut and oviduct.

4.4 A potential role of AeInx7 in epithelial and neural function

AeInx7 mRNA is expressed throughout the alimentary canal (midgut, hindgut and

Malpighian tubules) of both sexes (Figure 13). This expression pattern could indicate that AeInx7 is important for coordinating the respective digestive and/or excretory functions that these tissues are responsible for, such as the diuretic processes of the

Malpighian tubules, the release of digestive enzymes into the midgut, and the reabsorption of ions and water by the hindgut. AeInx7 mRNA is also expressed in the adult head and carcass (Figure 14), which may indicate an important role of this innexin in the nervous system. Consistent with this notion, in D. melanogaster, Inx7 is required for proper nervous system development during embryogenesis, and in memory formation by adult flies (Ostrowski et al. 2008; Wu et al. 2011).

4.5 The molecular potential for homomeric/heteromeric hemichannels and homotypic/ heterotypic gap junctions

The molecular expression data of Figures 13-14 also reveals the potential types of GJ that can be formed in specific tissues. For example, in the ovaries, the potential is high for the presence of homotypic and heterotypic GJ consisting of homomeric and/or heteromeric hemichannels, because this tissue expresses mRNAs of AeInx1, AeInx2,

AeInx3, AeInx4, and AeInx8 (Figure 14). Likewise, the expression of no less than four innexin genes in each of the adult tissues examined suggests that the molecular composition of GJ in mosquitoes is complex. Thus, a highly integrative set of technical approaches that includes cellular localization of the encoded proteins using antibodies,

84 functional characterization of the encoded proteins using a heterologous expression system (e.g., Xenopus oocytes), and knockdown of the mRNAs using RNA interference will likely be required to elucidate the precise functional roles of each innexin gene.

4.6 Cloned innexin cDNAs

This study is the first to clone innexin cDNAs and identify splice variants in the

Malpighian tubules of any insect. For both AeInx1 and AeInx3, the splice variation results in novel predicted proteins that differ at the extreme end of the cytosolic COOH- terminal domains. This finding is notable, because in the connexins of vertebrates, the

COOH-terminal domain appears to play important roles in membrane trafficking and voltage gating of GJ (L. Bao et al. 2007; X. Bao et al. 2004; R. G. Johnson et al. 2012).

Thus, the alterations to the amino-acid sequences of AeInx1 and AeInx3 caused by alternative splicing may affect the surface membrane expression and activity of the encoded proteins. Alternate splicing has also been documented in the innexins of D. melanogaster (DmInx). In DmInx2, splicing occurs outside of the ORF in the 5’-UTR and thus does not change the predicted peptide sequence (L. A. Stebbings et al. 2000).

However, in DmInx8, splicing affects the ORF leading to polypeptides with differing sequences of their NH2-terminal domains, and these variants are expressed differentially in separate tissues (Crompton et al. 1995).

Both AeInx3 and AeInx7 are predicted to have glycosylation sites in their second and first extracellular loops, respectively. This finding is interesting because in pannexins

(the innexin homologs of vertebrates) the second extracellular loop is glycosylated, which may prevent the formation of GJ, and instead lead to functional hemichannels in the plasma membrane (Scemes et al. 2009). Pannexin hemichannels are thought to play roles in ATP signaling, as well as in general cation transport (Scemes and Spray 2012;

Scemes et al. 2009). Thus, AeInx3 and AeInx7 have the molecular potential to become glycosylated, which may allow them to take on other functional roles, besides that of GJ connectivity. Moreover, at least for AeInx3, Western blots in adult females reveal the appearance of multiple immunoreactive bands greater than the expected size (Figure 7), which is consistent the presence of glycosylated forms.

Nearly all of the cloned innexins were found to have at least one potential phosphorylation site, which suggests further putative regulatory mechanisms of innexin function in A. aegypti. The predicted AeInx1-A protein has only one predicted phosphorylation site in the COOH-terminus, and this site is lost in the predicted AeInx1-

B protein as a result of alternative splicing. The phosphorylation of connexins can alter channel properties such as permeability and regulate protein trafficking (X. Bao et al.

2007; Lampe and Lau 2004). Thus, there is potential for the two AeInx1 proteins to have differential intercellular transport and/or trafficking properties.

4.7 AeInx3 localization

Immunolabeling experiments in the alimentary canal with the anti-AeInx3 antibody revealed trends in AeInx3 immunoreactivity that mirror those of AeInx3 mRNA

86 expression, suggesting that AeInx3 protein levels follow mRNA levels. The immunoreactivity of AeInx3 was strongest in the hindgut, where it localized to the intercellular regions of epithelial cells in the ileum and rectum. This finding suggests that

AeInx3 may play a role in coordinating the functions of the hindgut epithelial cells.

The immunoreactivity was next strongest in the Malpighian tubules, where it was prominent in principal cells of the proximal and distal segments. The labeling occurred at intercellular boundaries between principal cells and within discrete intracellular compartments of principal cells (perhaps in vesicles). The intercellular labeling presents strong evidence for a role of AeInx3 in mediating the known electrical coupling of principal cells (Masia et al. 2000; Weng et al. 2008). We did not find direct evidence for the expression of AeInx3 immunoreactivity in stellate cells, but in the distal segment we observed immunolabeling in principal cells that neighbor stellate cells

(Figure 18B, arrow). Thus, AeInx3 in principal cells has the potential to dock with opposing innexins in stellate cells and form heterotypic GJ. Additional experiments will be necessary to determine the identity of such innexin(s) in stellate cells, which would help support the hypothesis that stellate and principal cells are functionally coupled (K

W Beyenbach and Piermarini 2011; Piermarini et al. 2010; Piermarini and Calkins 2014).

The immunoreactivity was weakest in the midgut, where it was only found in the tracheal tubes attached to the gut, but not in the epithelium or surrounding muscle.

The weak immunoreactivity is consistent with the weak expression of AeInx3 mRNA

87 detected by RT-PCR and suggests that the GJ of the midgut epithelia cells may have distinct functional properties from those of the hindgut and Malpighian tubules.

4.8 Summary

The present study sheds new light on the molecular expression of innexins and their potential functional roles in mosquitoes. We demonstrate patterns of life stage- and tissue-specific expression for six innexins of A. aegypti and clone the cDNAs for four of these innexins, including two splice variants. We also characterize the localization of

AeInx3 throughout the alimentary canal of adult females. Taken together, the data allow for the generation of hypotheses on the putative functions of these GJ in vivo, which will require testing in future studies.

5. Acknowledgements

We thank Dr. Matthew Rouhier (OSU) and Ms. Nuris Acosta (OSU) their assistance in the laboratory and rearing mosquitoes, respectively. We also thank Dr. David L. Denlinger

(OSU) and Dr. Omprakash Mittapalli (OSU) for their critical reading of this manuscript.

Funded by NIH grant R03DK090186 to PMP.

Chapter 4: Gap junctions and the mosquito blood meal

Abstract

Mosquitoes are vectors of pathogens that cause diseases of medical and veterinary importance. Female mosquitoes transmit these pathogens while taking a blood meal, which most species require to produce eggs. The period after a blood meal is a time of extreme physiological change that requires rapid coordination of specific tissues. Gap junctions (GJ) are intercellular channels that aid in the coordination of cells within tissues via the direct transfer of certain small molecules and ions between cells. Evolutionarily distinct groups of proteins form the gap junctions of vertebrate and invertebrate animals

(connexins and innexins, respectively). Aedes aegypti mosquitoes possess six genes encoding innexins: inx1, inx2, inx3, inx4, inx7, and inx8. The goal of this study was to identify potential roles of gap junctions in the physiology of mosquitoes within 24 h after a blood meal by using qPCR to quantify changes in innexin mRNA expression in adult females at 3 h and 24 h post-blood meal (PBM) relative to non-blood-fed controls. We found that at 24 h PBM, expression levels of several innexin mRNAs were altered, and that one innexin in particular, inx2, was the most highly up-regulated innexin in key tissues associated with blood-meal digestion and egg production (i.e., the midgut and

89 ovaries, respectively). However, despite knocking down inx2 mRNA levels by over 75% via

RNA interference, we found no significant effect on fecundity. Altogether, our results suggest that a blood meal influences the molecular expression of innexins in mosquitoes, but their specific physiological roles remain to be elucidated.

1. Introduction

Mosquitoes transmit many pathogens that cause deadly and debilitating diseases.

The yellow fever mosquito, Aedes aegypti, is the primary vector of chikungunya, dengue, yellow fever, and Zika viruses. These pathogens are transmitted when an infected female mosquito feeds on the blood of a vertebrate host; females require blood in order to produce eggs. Following consumption of a blood meal, the mosquito undergoes a complex series of physiological changes that require the rapid endocrine coordination of multiple tissues (Hansen et al. 2014). Intercellular channels known as gap junctions allow for endocrine signals to be rapidly shared among adjacent cells within a tissue by mediating the direct transport of ions, small molecules, and second messengers between cells (P Phelan and Starich 2001).

Gap junctions are comprised of proteins known as connexins in vertebrates and innexins in invertebrates; they are evolutionarily distinct proteins that have convergently evolved to share similar structure and function (P Phelan et al. 1996; Pauline Phelan et al.

1998). Some of the broad functional roles that innexin and connexin channels contribute to include embryonic development, post-embryonic development, immune response,

90 and reproduction (Bosco, Haefliger, and Meda 2011; Giuliani et al. 2013; Hasegawa and

Turnbull 2014; Holcroft et al. 2013; Li et al. 2014; Tazuke et al. 2002). However, the physiological roles of gap junctions in mosquitoes have only recently begun to emerge.

Prior work in Ae. aegypti has 1) demonstrated that innexin mRNA expression is life stage and tissue dependent, 2) implicated gap junctions in the physiology of renal

(Malpighian) tubules and the diuretic capacity of adult female mosquitoes, and 3) shown that gap junctions are critical for the survival of larval and adult female mosquitoes

(Calkins et al. 2015; Calkins and Piermarini 2015; Piermarini and Calkins 2014; Weng et al.

2008). In Anopheles gambiae, inx4 (a.k.a. zero population growth or zpg) is required for proper male gonad formation, while inx1 is necessary for immune response to invading

Plasmodium parasites (Li et al. 2014; Magnusson et al. 2011). As such, gap junctions appear to play vital roles in mosquito biology.

Here we aimed to test the hypothesis that gap junctions contribute to the post- blood meal physiology of Ae. aegypti by comparing the expression of innexin mRNAs in tissues involved with blood digestion (i.e., midgut), excretion of blood meal metabolites

(i.e., Malpighian tubules), and egg development (i.e., fat body and ovaries), between non- blood fed and blood fed mosquitoes. We found that at 24 h post-blood meal, the expression of at least one innexin mRNA was altered in the midgut, Malpighian tubules, ovaries, and whole mosquito. Although inx2 was the most highly up-regulated innexin in the midgut and ovaries, knockdown of inx2 mRNA levels via RNAi yielded no effect on fecundity.

2. Methods

2.1 Mosquito rearing

Eggs of Ae. aegypti were obtained through the Malaria Research and Reference

Reagent Resource Center (MR4) as part of the BEI Resources Repository (Liverpool strain;

LVP-IB12 F19, deposited by M.Q. Benedict). Mosquitoes were reared to adults and the colony was maintained as described in Piermarini et al. (2011). In brief, larvae and adults were held in an environmental chamber set at 28C and 80% relative humidity with a 12 h:12 h light:dark cycle. Larvae were reared in plastic trays in distilled water and fed ground

TetraMin tropical fish flakes (Tetra Spectrum Brands, Blacksburg, VA). Adults were housed in 0.5 m3 cages with access to 10% sucrose, and fed heparinized rabbit blood (Hemostat

Laboratories, Dixon, CA) through a membrane feeder (Hemotek, Blackburn, UK) to produce eggs.

2.2 Blood feeding

The mosquitoes to be blood fed were adult females 3-7 days post eclosion.

Twenty-four hours prior to blood feeding, sucrose was removed from the mosquito cages.

Mosquitoes were then offered heparinized rabbit blood (Hemostat Laboratories) for 60 min using an artificial membrane feeding system (Hemotek). Control mosquitoes (non- blood fed) were treated similarly and provided access to 10% sucrose instead of blood for

60 min. After blood feeding, both groups were provided with 10% sucrose. Mosquitoes for tissue level qPCR analysis were dissected at either 3 h or 24 h post-blood meal.

2.3 Dissection, RNA extraction and cDNA synthesis

Dissections, RNA extraction, and cDNA synthesis were performed as described in

Calkins et al. (2015). In brief, mosquitoes were anesthetized on ice before being dissected in mosquito Ringer solution (consisting of 150 mM NaCl, 3.4 mM KCl, 1.7 mM CaCl2, 1.8 mM NaHCO3, 1 mM MgCl2, 5 mM Glucose, 25 mM HEPES; pH 7.1). Tissues were isolated, transferred to 1.5 ml micro-centrifuge tubes (Thermo Fisher Scientific, Waltham, MA), and preserved in TRIzol® reagent at -80°C until utilized in RNA isolation. Total RNA was isolated using the method of Chomczynski and Sacchi (1987) and quantified using a

NanoDrop 2000 spectrophotometer (Thermo Scientific, Wilmington, DE). cDNA libraries were synthesized using 4 µg of total RNA and the GoScriptTM Reverse Transcriptase system with random primers (Promega, Madison, WI), following manufacturer’s protocols. cDNA libraries were stored at -20° C until needed for qPCR.

2.4 qPCR

qPCR was performed as described in Calkins and Piermarini (2015). In brief, reactions were performed in triplicate with each reaction consisting of 5 µl of GoTaq® qPCR Master Mix, 400 nM forward and reverse primers, 40 ng cDNA and nuclease free water (total volume = 10 µl). Primers for each innexin and our reference gene (Ribosomal

93 protein S7, RPS7) were used as in Calkins and Piermarini (2015; Table 9). The reactions were then subjected to the following thermocycling protocol using a C1000/CFX96 real time system (Bio-Rad Laboratories, Hercules, CA): initial denaturation of 95 °C (3 min), followed by 39 cycles of 95 °C (10 sec), and 58 °C (30 sec), ending with a melt curve cycle. qPCR results were analyzed using the ΔCt method (Silver et al. 2006; Song et al. 2012) and expressed as relative gene expression.

Table 9: qPCR primer pairs. Each set of innexin primers was determined to be specific through melt curve analysis and DNA sequencing of PCR products.

qPCR Forward qPCR Reverse Inx1 CACCGATAGTGCCGTATTCC CCGACATATTGTGTGGCAGT Inx2 GGAGATCCTATGGCACGAG ACGGTAGCACACAGAGTCC T A Inx3 TCGTTCGGTTACTTCATCTG GCGATTCTCCTGATCCATGT C C Inx4 TTCTGTTGGACACTGGGAA CCATGTGCGTTCCTATTTCG C Inx7 TGGGTCCCGTTTGTGTTATT CCATACGAAGACCATCCAC A Inx8 GACTGCGTTCACACGAAAG GGGTACTTCGCTACCGACTT A T RPS7 CTTTGATGTGCGAGTGAAC CATCTCCAACTCCAGGATAG AC C

2.5 dsRNA synthesis and injections

dsRNA synthesis and injections were performed as described in Calkins and

Piermarini (2015). In brief, primers for dsRNA template synthesis were designed to amplify 300-500 bp gene segments (Calkins and Piermarini 2015; Table 10) to be utilized 94 in the T7 MEGAscript® dsRNA synthesis kit (Thermo Fisher Scientific). dsRNA was synthesized following the manufacture’s protocols and stored at -80°C.

Table 10: dsRNA template synthesis primers. Each primer set consists of an innexin specific region for amplification of the target gene from plasmid, and the T7 promoter sequence (TAATACGACTCACTATAGGGAGA).

dsRNA Template Forward dsRNA Template Reverse Inx2 TAATACGACTCACTATAGGGA TAATACGACTCACTATAGGGA GATTTGGCGTTTGAAAAGTGT GAATACTCCCGGCTGAGCAAT G A eGFP TAATACGACTCACTATAGGGA TAATACGACTCACTATAGGGT CGTAAACGGCCACAAGTT TGGGGTCTTTGCTCAGG

On the day of an injection, dsRNA was diluted to 1 µg/µl in 0.5X PBS solution (5.95 mM phosphates, 68.5 mM sodium chloride and 1.35 mM potassium chloride; pH 7.5;

Fisher Scientific, Fairlawn, NJ). Eighty mosquitoes were injected with 1 µg of either inx2 dsRNA or eGFP dsRNA and returned to rearing conditions with access to 10% sucrose.

After three days, three mosquitoes were removed from each treatment for knockdown assessment via qPCR and the remaining mosquitoes were utilized in fecundity assays (see section 2.6). Non-blood fed mosquitoes were utilized in knock down analysis for phenotype assessment as knockdown was not significantly different between blood fed and non-blood fed mosquitoes (Figure 21). For tissue level knockdown analysis, forty mosquitoes were injected with dsRNA for either inx2 or eGFP and dissected three days post injection.

Figure 21: Knockdown of inx2 in blood fed and non-blood fed mosquitoes. Bars indicate means ± SEM, n=3. Letters represent statistical differences as determined by a two-way ANOVA. Knockdown of inx2 is not significantly different between blood fed and non- blood fed mosquitoes, 24 hours post blood meal.

2.6 Fecundity and viability assays

Three days after dsRNA injection, mosquitoes were offered a blood meal (see section 2.2); those with no visible blood in their abdomens were excluded from the assays. Twenty-four hours later, the blood-fed mosquitoes were transferred to individual 96 egg-laying tubes for the fecundity assay. The egg-laying tubes consisted of a cylindrical glass tube (21 X 70 mm; Fisher Scientific, Pittsburg, PA) with a piece of coffee filter

(Melitta USA, Clearwater, FL) cut to fit the bottom of the tube. The filter was wetted with

150 µl of ddH2O (Milli-Q® filtered water, Merck KGaA, Darmstadt, Germany) and the open end was plugged with a cotton ball. The mosquitoes in their individual egg laying tubes were returned to rearing conditions for 48 h, and the number of eggs laid by each mosquito was counted.

After counting the eggs, filter papers were consolidated according to dsRNA treatment and allowed to dry in a rearing chamber for one week. Eggs were then hatched in ddH2O under vacuum and returned to rearing conditions for 24 h. The resulting larvae were immobilized through refrigeration before counting them under a dissection stereomicroscope (World Precision Instruments, Sarasota, FL/model PZMTIII-BS).

2.7 Data Analysis

All data were analyzed with GraphPad Prism 6 (GraphPad Software, La Jolla, CA).

Differences in innexin mRNA expression were analyzed via a one-way ANOVA (non-blood fed tissue analysis) or two-way ANOVA (blood fed vs. non-blood fed tissue analysis).

Significant gene knockdown via RNAi was determined by t-test. Differences in percent viability and in percent mosquitoes ovipositing were determined by one-way ANOVAs.

Difference in number of eggs laid was determined by Kruskal-Wallis test with Dunn’s multiple comparison test post hoc analysis.

3. Results

3.1 Gene expression analysis

For each of the six innexins in Ae. aegypti (inx1, inx2, inx3, inx4, inx7, and inx8), we compared mRNA expression among the Malpighian tubules, midgut, ovaries, and fat body, which was normalized to expression in the whole insect (WI). As shown in Figure

22, we found that: 1) inx1 is most abundant in the fat body (~2.8-times WI); 2) inx2 is ubiquitously expressed across the tissues examined (~1-2-times WI); 3) inx3 is most abundant in the Malpighian tubules (~2.7-times WI) and least abundant in the midgut

(~0.2-times WI); 4) inx4 is highly abundant in the ovaries (~6.5-times WI); 5) inx7 is highly abundant in the midgut (~10.6-times WI); and 6) inx8 is weakly expressed in all tissues examined (~0.3-times WI), but is highest in the ovaries (~0.9-times WI).

Figure 22: Tissue specific expression of innexins in non-blood fed females. Bars represent expression relative to whole insect (WI) expression levels. Values are means ± SEM. n=10 biological replicates. MT, Malpighian tubules; MG, midgut; OV, ovaries; FB, fat body. Letters represent differences as determined by a one-way ANOVA.

Next, we compared innexin mRNA expression between 1) blood fed and non- blood fed females and 2) tissues isolated from blood fed and non-blood fed females (i.e.,

Malpighian tubules, midgut, ovaries, and fat body). At 3 h post-blood meal (PBM), there were no significant differences in innexin mRNA levels between blood fed and non-blood fed mosquitoes (Figure 23). However, at 24 h PBM, at least one innexin was differentially expressed between blood fed and non-blood fed mosquitoes at the whole mosquito and tissue levels with the exception of the fat body (Figure 24). At the whole insect level, inx2, inx3, and inx4 were significantly up-regulated. At the tissue level, inx2 was significantly up-regulated in the Malpighian tubules, midgut, and ovaries, and inx3 was significantly up-regulated in the ovaries.

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Figure 23: Effects of a blood meal on innexin mRNA expression 3-h post-blood meal. White bars indicate non-blood fed control females and red bars indicate blood fed females. Bars indicate means ± SEM, n=5. Abbreviations are as in Figure 22. No differences were found in innexin expression between blood fed and non-blood fed controls as determined by a two-way ANOVA.

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Figure 24: Effects of a blood meal on innexin mRNA expression 24 h post-blood meal. White bars indicate non-blood fed control females and red bars indicate blood fed females. Bars indicate means ± SEM, n=5. Abbreviations are as in Figure 22. Asterisks indicate differences within a gene between blood fed and non-blood fed controls as determined by a two-way ANOVA and Newman-Keuls multiple comparison.

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3.2 RNAi:

Given that inx2 was up-regulated in the whole insect, Malpighian tubules, midgut, and ovaries at 24 h PBM, it was selected for RNAi experiments. Injecting 1 µg of inx2 dsRNA into adult females resulted in a 75.7 ± 3.9% reduction in mRNA expression (relative to controls injected with eGFP dsRNA) at the whole insect level within 3 days with no significant changes in expression of the other innexin mRNAs (Figure 25A). At the tissue level, knockdown of inx2 was similar in the midgut (94.8 ± 0.5%), ovaries (88.5 ± 0.2%), and fat body (91.0 ± 0.7%), while knockdown of inx2 in the Malpighian tubules was significantly weaker (44.9 ± 10.1%) (Figure 25B).

Figure 25: RNAi-induced knockdown of inx2 mRNA expression. A) Whole insect innexin mRNA expression 3 days post injection of inx2 dsRNA (percent relative to eGFP dsRNA- injected controls). Asterisk indicates significant difference from eGFP dsRNA-injected mosquitoes as determined by a T-test. Bars represent means ± SEM, n=7. B) Tissue-level innexin mRNA expression 3 days post injection of inx2 dsRNA. Abbreviations are as in Figure 22. Bars represent means ± SEM, n=3. Letters represent differences as determined by one-way ANOVA and Newman-Keuls multiple comparison.

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Despite the effective and specific knockdown, inx2 dsRNA injection was without significant effects on the median number of eggs laid (55 eggs per mosquito) compared to uninjected controls (60 eggs per mosquito; Figure 26A). However, mosquitoes injected with eGFP dsRNA laid a significantly fewer median number of eggs (40 eggs per mosquito) than both the inx2 dsRNA-injected and uninjected mosquitoes (Figure 26A). There was neither a significant difference in the viability of eggs among the groups (eGFP = 56.7 ±

4.6%; inx2 = 45.4 ±-6.8%; uninjected = 64.1 ± 4.3%; Figure 26B), nor the percentage of mosquitoes that oviposited among our treatments (eGFP = 67.4 ± 4.5%, inx2 = 76.5 ±

3.9%, uninjected = 72.3 ± 15.7%; Figure 26C).

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Figure 26: Effects of inx2 knockdown on fecundity of adult female mosquitoes. A) Horizontal bar represents mean percentage of mosquitoes ovipositing during the assay. Points are representative of independent replicates (n=7 for eGFP, 7 for inx2, and 3 for untreated). B) Horizontal bar represents median number of eggs laid per mosquito. Columns are labeled by dsRNA treatment. Points are representative of individual mosquitoes (n=231 for eGFP, 235 for inx2, and 119 for untreated). Letters represent differences as determined by a non-parametric one-way ANOVA and Dunn’s multiple comparison. C) Horizontal bar represents mean percentage of eggs hatching into viable larvae. Points are representative of independent replicates (n=7 for eGFP, 7 for inx2, and 3 for untreated).

4. Discussion

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Our study provides the first quantitative characterization of innexin mRNA expression in adult female Ae. aegypti following a blood meal, with a specific focus on the

Malpighian tubules, midgut, ovaries, and fat body. We also used RNAi to investigate a potential role of inx2 in fecundity given its dramatic up-regulation in the midgut and ovaries 24 h post-blood meal. Taken together, the data from this study provide insights into the molecular expression of innexin genes in adult female Ae. aegypti mosquitoes and identify a potential role of inx2 in the post blood meal physiology of Ae. aegypti, albeit one that is apparently unrelated to fecundity.

4.1 The molecular expression of innexins in isolated tissues of non-blood fed mosquitoes

The quantitative patterns of innexin mRNA expression that we characterized in the tissues of non-blood fed mosquitoes (Fig. 22) were generally consistent with those found in our previous qualitative RT-PCR study of innexin mRNA expression (Calkins et al

2015). One notable exception was the expression pattern of inx7. That is, we previously considered inx7 mRNA expression to be abundant in both the Malpighian tubules and midgut of adult females (Calkins et al. 2015). However, in the present study we determined that inx7 mRNA levels in the Malpighian tubules were significantly lower than those in the midgut (Fig. 22). Notably, inx7 mRNA was only abundantly expressed in the midgut, suggesting that it might be involved in digestion and/or nutrient absorption.

Reverse genetic experiments will be required to elucidate its functional role in the midgut.

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The present study is the first to characterize innexin mRNA expression in the fat body of Ae. aegypti. A previous transcriptomic analysis of the fat body, detected innexin mRNA expression (Price et al. 2011), but their low abundance compared to other transcripts makes quantitative comparisons difficult. Using qPCR, we found that inx1 mRNA was particularly abundant in the fat body, whereas inx2, inx3, and inx8 mRNAs were expressed at levels comparable to those found in other tissues (Figure 22). Neither inx4 nor inx7 mRNAs were detectable in the fat body. The molecular expression of innexins detected in the present study, paired with the ultrastructural evidence for gap- junctional plaques in the fat body of mosquitoes (Dean, Collins, and Locke 1985), suggests that innexins may be involved in key physiological roles that the fat body plays, such as lipid storage and utilization, vitellogenesis, and/or immune response (Arrese and

Soulages 2010; Hansen et al. 2014; Tsakas and Marmaras 2010). Reverse genetic experiments will be required to elucidate specific functional roles of innexins in the fat body.

4.2 A blood meal influences innexin mRNA expression

The up-regulation of inx2, inx3, and inx4 at 24 h PBM in the whole insect can in part be explained by an up-regulation of inx2 in the Malpighian tubules, midgut, and ovaries, and inx3 in the ovaries (Figure 24). An up-regulation of inx4 was not found in any of the tissues examined in the present study (Figure 24). Based on our previous qualitative

RT-PCR study, inx4 mRNA in adult female Ae. aegypti was only detectable in the ovaries

107 and carcass (Calkins et al. 2015); the carcass includes the thoracic and abdominal body walls after removing the head, alimentary canal, and gonads. Given the lack of differential expression after blood feeding in the fat body (Fig. 24), which includes the abdominal body wall, it would appear that the thorax is the most likely site of inx4 up-regulation post-blood meal. The specific role that inx4 may be playing in the thorax is unclear, but it could be related to changes in the activity of the thoracic flight musculature post-blood meal (Jones and Gubbins 1978; Marinotti et al. 2005).

4.3 Potential roles of inx2 in the physiology of mosquitoes PBM

As mentioned above, at 24 h PBM, inx2 mRNAs were up-regulated in 3 distinct tissues that play key roles in blood meal processing and egg development: the midgut,

Malpighian tubules, and ovaries. After a blood meal, the midgut contributes to digestion, nutrient absorption, immune responses, and xenobiotic detoxification, while the

Malpighian tubules contribute to diuresis as well as xenobiotic and metabolite detoxification/excretion (Esquivel, Cassone, and Piermarini 2014, 2016; Holt et al. 2002;

Sanders et al. 2013). Additionally, the ovaries undergo vitellogenesis and dramatically increase in size before eggs are oviposited (Koller and Raikhel 1991; Raikhel 1992). Thus, these tissues undergo profound physiological changes after the mosquito ingests a blood meal. The consistent up-regulation of inx2 in all three of these tissues at 24 h PBM suggests that inx2-mediated intercellular communication may contribute to the regulation of the physiological activities in these tissues. Alternatively, inx2 could be

108 acting as a hemichannel in the plasma membranes of these tissues to release signaling molecules that contribute to paracrine/endocrine communication after a blood meal, as suggested by Li et al. (2014) for inx1 in the midgut of An. gambiae.

We hypothesized that knockdown of inx2 would disrupt digestion, excretion, and/or oogenesis after a blood meal, thereby resulting in a reduction of fecundity.

However, despite knocking down inx2 mRNA levels by ~75% in whole mosquitoes, ~45% in the Malpighian tubules, and ~90% in midgut and ovaries (Fig. 25), we found no reduction in the percentage of mosquitoes that oviposited, the number of eggs laid per mosquito, or the viability of the eggs in inx2 dsRNA-injected females compared to uninjected mosquitoes (Fig. 26). While this may suggest inx2 is not directly related to fecundity, without protein level analysis, we cannot rule out the possibility that protein levels of inx2 were unaffected by the mRNA knockdown, thereby leading to a lack of phenotype. Alternatively, inx2 may be playing an important physiological role outside of fecundity after the blood meal. For example, in An. gambiae, inx1 is necessary for singalling in the upregulation of Tep1, a gene involved in insect innate immune response.

Intriguingly, eGFP dsRNA-injected mosquitoes laid fewer eggs than the inx2 dsRNA- injected and uninjected mosquitoes (Fig. 26), suggesting a potential consequence of eGFP dsRNA injection on fecundity. Although the mechanism by which this would occur is unclear, in the honeybee, Apis mellifera, injection of GFP dsRNA results in differential expression of ~10% of the insect’s transcriptome, including an up-regulation of immune genes (Nunes et al. 2013). Moreover, in D. melanogaster, RNAi mechanisms are part of

109 the immune response to viral dsRNA infection (Zambon, Vakharia, and Wu 2006). Thus, it is possible that injection of eGFP dsRNA in Ae. aegypti elicits an anti-viral immune response that diverts energetic resources from oogenesis. It remains to be elucidated whether inx2 is involved with an anti-viral response, which could explain the higher fecundity in the inx2-dsRNA injected mosquitoes.

5. Conclusions

The present study is the first to quantify effects of a blood meal on molecular expression of innexins in Ae. aegypti. While we found inx2 to be highly up-regulated in both the midgut and ovaries following a blood meal, RNAi of inx2 yielded no reduction in fecundity. While this may suggest a role of inx2 outside of reproduction, experiments assessing protein levels in the mosquito are necessary to verify that mRNA knockdown is having the same effect on protein levels. This quantitative study also confirms the general qualitative trends in innexin expression demonstrated previously (Calkins et al 2015).

Taken together, the results from this study provide molecular evidence that innexins are potentially important in the physiology of Ae. aegypti after a blood meal, but their specific functional roles remain to be elucidated.

6. Acknowledgements

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We thank Dr. David Denlinger (OSU) for his critical review of this manuscript and

Ms. Nuris Acosta (OSU) and Ms. Edna Alfaro (OSU) for their assistance in mosquito rearing. Funding for this study was provided by grants to 1) PMP from the NIH

(R03DK090186) and Mosquito Research Foundation (2014–03), and 2) TLC from the

OARDC SEEDS program (Grant# 2014–078; oardc.osu.edu/seeds) and Grants-in-Aid of research from both The Ohio State University and national chapters of the Sigma-Xi scientific research society. State and Federal funds appropriated to the OARDC of the

Ohio State University also supported the study.

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Chapter 5: The Contractile Nature of the Mosquito Crop

Abstract

The crop is a diverticulum of the esophagus and a food storage organ conserved among flies (Order: Diptera). The crop must pump its stored contents back into the alimentary canal for digestion and absorption by the animal. The pumping is mediated by peristaltic contractions of the crop musculature. In adult female mosquitoes, the crop selectively stores sugar solutions (e.g., nectar); proteinaceous blood meals bypass the crop and are directly transferred to the midgut for digestion. The mechanisms that mediate and regulate crop contractions have never been investigated in mosquitoes.

However, these mechanisms are relatively well described in other flies, such as the blow fly (Phormia regina). Here we characterized the contractile nature of the mosquito crop, and investigated the potential involvement of gap junctions in its contraction. To accomplish this, we measured contraction rates of mosquito crops in vitro, and utilized qPCR and immunohistochemistry to characterize the expression of gap junctional proteins (i.e., innexins). We found that the mosquito crop is under similar physiological controls as other flies, with serotonin increasing crop contractions and a dromyosupressin mimic, benzethonium chloride, decreasing contractions. We also implicated gap junctions 112 in crop contraction, with the gap junction inhibitor carbenoxolone reducing crop contraction. Furthermore, we localized protein immunoreactivity of inx2 to muscle cells and inx3 to epithelial cells. Finally, using qPCR we identified Inx2 as the most highly expressed innexin in the crop. Taking our findings together, we developed a model of the signaling pathways for crop muscle contraction.

1. Introduction

The alimentary canal of adult dipteran insects possesses a diverticulum of the foregut referred to as the crop or ventral diverticulum. The crop consists of 4 main structures: 1) luminal cuticle, 2) a simple epithelium, 3) a network of muscles, and 4) nerves that derive from the corpus cardiacum (Stoffolano and Haselton 2013). Food storage is the quintessential physiological function of the crop. That is, it receives imbibed liquids (e.g., nectar) and stores them for later digestion and absorption by the midgut.

The release of food from the crop to the midgut is mediated by peristaltic contractions of the crop musculature, which are regulated by an array of neural and neuroendocrine mechanisms and physiological factors.

In the blow fly (Phormia regina), extracellular Ca2+ is essential for crop muscle contraction, and hemolymph osmolality can influence the rate of contraction (Gelperin

1966; Liscia et al. 2012; Solari et al. 2013). Moreover, the crop contraction rates in both the blow fly and house fly (Musca domestica) generally increase as crop volume increases

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(Holling 1976; Stoffolano, Patel, and Tran 2014). In addition to hemolymph factors and crop volume, myo-suppressive neuropeptides, such as dromyosuppressin (DMS), and myo-active neurotransmitters, such as serotonin, contribute to the regulation of crop contraction rates in the blow fly (Liscia et al. 2012; Stoffolano, Danai, and Chambers

2013). The crop of the fruit fly (Drosophila melanogaster), is similarly under neuronal control, with DMS, drosulfakinin, and FMRFamide all reducing contraction rates

(Duttlinger, Berry, and Nichols 2002; Palmer et al. 2007). Finally, mutations in the fruit fly can also influence crop contraction, with drop-dead knockouts showing increased crop contraction rates, but a failure of crop emptying (Peller et al. 2009).

In mosquitoes, the crop is the primary storage organ for imbibed sugar (e.g., nectar) before it is pumped into the midgut for digestion. In contrast, blood, which is only ingested by adult females, bypasses the crop and is received directly by the midgut for immediate digestion (Day 1954). Sugar feeding has been implicated in the longevity and fecundity of mosquitoes, both of which can directly influence vectorial capacity (Foster

1995). Immunochemical studies in the mosquito crop have identified neurons that contain FMRFamide, small cardioactive peptide b, and serotonin (Moffett and Moffett

2005), but the physiological mechanisms that regulate its contractile activity have not been previously investigated. Here we provide the first characterization of the basic mechanisms that contribute to the regulation of crop contractions in adult female Aedes aegypti, an important vector of arboviruses that cause Zika, dengue, chikungunya, and yellow fevers in humans. Moreover, we test the hypothesis that gap junctions are

114 involved with the peristaltic contractions of the crop. Gap junctions are intercellular channels that play key roles in the coordination of neuromuscular function and/or electrical coupling of cells in a wide range of animal tissues, including the vertebrate heart and mosquito Malpighian tubules (Degroot et al. 2003; Piermarini and Calkins 2014; Weng et al. 2008), but their role and presence in the insect crop have not been previously examined.

Utilizing an in vitro assay, we show that the isolated mosquito crop spontaneously contracts in Ringer solution for at least one hour and its contractions are dependent upon the presence of extracellular Ca2+. Moreover, the contractions are potently stimulated by serotonin (5 hydroytryptamine, 5-HT), inhibited by a chemical agonist of the DMS receptor (benzethonium chloride), and unaffected by a kinin (aedeskinin III). The agonistic effects of serotonin are mimicked by a membrane-permeable analog of cyclic adenosine monophosphate (cAMP), and inhibited by an inhibitor of protein kinase A, consistent with signaling via a GPCR that increases cAMP and activates PKA. Notably, for the first time in any insect, we provide pharmacological evidence that gap junctions are required for crop contractions, and molecular and immunochemical evidence for the expression of gap junctional proteins (innexins) in muscle and epithelial cells of the crop.

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2. Methods

2.1 Mosquito rearing

To establish our colony, Aedes aegypti mosquitoes were obtained as eggs through the Malaria Research and Reference Reagent Resource Center (MR4) as part of the BEI

Resources Repository (Liverpool strain; LVP-IB12 F19, deposited by M.Q. Benedict). In brief, mosquitoes were reared in an environmental chamber set to 28°C and 80% relative humidity with a 12 h:12 h light:dark cycle, as described in (Piermarini et al. 2011). Larvae were fed daily with pulverized Tetramin flakes and adults were fed 10% sucrose ad libitum.

2.2 Crop dissection

At 3-10 days post-emergence, adult female mosquitoes were removed from rearing cages and immobilized on ice. Only females with visibly distended abdomens (i.e., obvious signs of recent sugar feeding) were used. After removing the legs, the body was submerged in mosquito Ringer solution (150 mM NaCl, 3.4 mM KCl, 1.7 mM CaCl2, 1.8 mM NaHCO3, 1 mM MgCl2, 5 mM Glucose, and 25 mM HEPES; pH 7.1; 331 mOsm/kg) at room temperature. The head was removed with forceps (Dumont #5; Fine Science Tools,

Inc., Foster City, CA) under Ringer solution and the thorax was gently teased away from the abdomen, exposing the crop and its attachment to the foregut of the alimentary

116 canal. The foregut and anterior midgut were compressed with forceps anteriorly and posteriorly to the crop, allowing it to be isolated from the alimentary canal.

2.3 In vitro contraction assays

To measure crop contraction rates, we used an in vitro assay similar to that used in the blow fly by Haselton et al. (2006). In brief, crops for in vitro contraction assays were transferred by glass pipette directly into a single well of a 96-well microtiter plate. (USA

Scientific) containing 100 µl of mosquito Ringer solution. The contraction rates of crops were stable between 5 and 20 minutes post-transfer (Figure 27A). Thus, all measurements and experiments were performed within this 15 minute window.

To test putative inhibitors and agonists of crop contraction rate, each crop served as its own control. In brief, after transferring the crop to a well, it was allowed 5 min to recover and then contractions were counted at 5, 7, and 9 mins post-transfer. These rates were averaged together and referred to as the ‘control’ contraction rate. At 10 min post- transfer, 1 µl of Ringer solution was removed from the well and replaced with 1 µl of a treatment solution, thereby diluting the treatment solution 100-times (see below). After gentle mixing via pipetting, the number of contractions per minute was counted again at

12, 14, and 16 min post-transfer. These rates were averaged together and referred to as the ‘treatment’ contraction rate.

The treatment solutions added to the wells consisted of serotonin at various concentrations, 100 mM 8-Bromo-cAMP sodium salt, 2.5 mM benzethonium chloride (a

117 dromyosupressin mimic), or carbenoxolone at various concentrations (CBX; an inhibitor of gap junctions). In some experiments, we tested the effects of serotonin (10 nM final bath concentration) on crops that were pre-treated with CBX (250 µM final bath concentration) or H-89 (an inhibitor of PKA, 10 µM final bath concentration). For these experiments, 1) contraction rates were counted at 5 and 7 min post-transfer for the control period, 2) CBX or H-89 (resuspended in DMSO; final bath concentration 1%) was added at 8 min post-transfer and contractions rates were counted at 10 and 12 min post- transfer, and 3) serotonin was added at 13 min post-transfer and contraction rates were counted at 15 and 17 min post-transfer. To determine the effects of extracellular Ca2+ on contraction rates, the crops were dissected—and the control contraction rates were measured—in a Ca2+-free Ringer solution; the treatment solution consisted of 170 mM

CaCl2.

2.4 RNA isolation and qPCR

Following their isolation from the mosquito, crops for gene expression analysis were transferred with forceps directly into TRIzol® reagent (Life Technologies, Carlsbad,

CA) on ice. RNA isolation and qPCR were performed as described in Calkins et al. (2015).

In brief, RNA was isolated using the method of Chomczynski and Sacchi (1987). cDNA was synthesized using 1 µg of total RNA in the GoScript® Reverse Transcriptase kit (Promega,

Madison, WI). Primers for qPCR were as used in Calkins and Piermarini (2015). For a given sample, each reaction consisted of 10 μl: 5 μl of GoTaq Master Mix, 40 ng cDNA (0.2 µl of

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200 ng/µl cDNA), 1 µl of 4 µM primers, and 3.8 μl nuclease free water. Three technical replicates were performed for each sample. The reactions took place in 96-well unskirted, low profile plates (Bio-Rad Laboratories, Hercules, CA), sealed with TempPlate RT optical film (USA Scientific, Orlando, FL). The qPCR utilized a Bio-Rad C1000 thermocycler and

CFX96 real time system (Bio-Rad Laboratories) with the following protocol: initial denaturation of 95°C (3 min) followed by 40 cycles of 95°C (10 sec) and 58°C (30 sec), ending with a melt curve analysis.

2.5 Antibodies

The polyclonal rabbit anti-AeInx3 antibody was previously used in Calkins et al.

(2015). In brief, the antibody was produced and affinity purified against a COOH-terminal peptide of the predicted AeInx3 protein (L374EMAPIYPEIGKYGKDTA). The polyclonal rabbit anti-Bminx2 antibody was a gift from Dr. Ryoichi Yoshimura of the Kyoto Institute of

Technology, Department of Applied Biology. The peptide used to develop the affinity- purified antibody corresponds to the first extracellular loop of inx2 in Bombyx mori

(VGPHVEGQDEVKYHK; Fushiki, Yoshimura, and Endo 2010).

2.6 Immunohistochemistry (IHC)

IHC was performed as described in Calkins et al. (2015). In brief, crops were fixed for 45 min at room temperature in a phosphate-buffered saline (PBS) containing 4% paraformaldehyde, and then washed 3 times for 5 min each in PBS; the PBS consisted of

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11.9 mM phosphates, 137 mM sodium chloride, and 2.7 mM potassium chloride (pH 7.5).

The fixed crops were permeabilized with 0.1% Triton X-100 (Thermo Fisher Scientific) for

20 min (two 10 minute washes), washed 3 times in PBS with 0.1% Tween 20 (Thermo

Fisher Scientific; PBT) for 5 min each, and then blocked in 10% normal goat serum

(Thermo Fisher) supplemented with 20% casein (Vector Laboratories, Burlingame, CA) for

2 hours (all at room temperature). The crops were then incubated with one of the following anti-innexin primary antibodies overnight at 4° C: rabbit anti-AeInx3 (1:400 in

PBT) or rabbit-anti-BmInx2 (1:100 in PBT). After the overnight incubation, the crops were washed 3 times in PBT for 5 min each, and incubated with DyLight 488 Goat-anti-rabbit secondary (Thermo Scientific; 1:600) for 2 h, before washing again 3 times with PBT for 5 min each (all at room temperature). Nuclei and muscle fibers were counterstained respectively with DAPI (ThermoFisher Scientific) and phalloidin conjugated to DyLight 633

(ThermoFisher Scientific). Labeled crops were imaged using a Leica S5 confocal microscope at the Molecular and Cellular Imaging Center (MCIC) of the Ohio Agricultural

Research and Development Center (OARDC) in Wooster, OH.

2.7 Statistical analysis

GraphPad Prism 6 software (GraphPad Software Inc., La Jolla, CA) was used in all statistical analysis. Individual tests are described in the respective figure legends.

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3. Results

3.1 The spontaneous contractions of isolated crops are stable and Ca2+-dependent

When transferred to 100 µl of Ringer solution in a 96-well microtiter plate, the crops spontaneously contracted at a rate of ~4 contractions/min. Within 2 min, the frequency significantly increased to ~6 contractions/min, then slightly reduced to ~5 contractions per minute and remained stable over the next 15 min (Figure 27A). By 30 min post-transfer, the contraction rate significantly reduced to ~3 per minute. At 60 min post-transfer, the contraction rate was similar to that at 30 min. Thus, the crop remained viable in vitro for at least 60 min and the contraction rate was highly stable between 5 and 20 min post-transfer (Figure 27A). Notably, if the crops were dissected in—and transferred to—a Ca2+-free Ringer solution, the contraction rates were nominal (~1 per min) within 5-10 min post-transfer. However, when Ca2+ was added to the bath, raising the concentration to that of normal Ringer solution (1.7 mM), the frequency significantly increased to ~6 contractions per min (Figure 27B); i.e., statistically similar to the contraction rates of crops in normal Ringer solution 5-20 min post-transfer (Figure 27A).

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Figure 27: In vitro crop assay panel. A) Baseline crop contractions from dissection through 1 hour post transfer. Times within the red box (5, 10, 15, and 20 minutes) were not significantly different than one another as determined by a repeated measures ANOVA with Newman-Keuls post hoc analysis, n = 10, P < 0.05. Subsequent assays were conducted between 5-20 minutes post-transfer. B) Effects of Ca2+ on crop contraction rate. Points represent mean crop contraction and error bars indicate SEM (n = 7). The first three points are control time points (no extracellular Ca2+ present), the arrowhead indicates the point of Ca2+ addition to the Ringer solution, and the final three points indicate experimental time points (Ca2+ present). Control points are all statistically equivalent, as are experimental points, and all experimental points are significantly greater than the control points as determined by a repeated measures ANOVA and Newman-Keuls post hoc analysis, p < 0.05. C) Effects of serotonin on crop contraction rate. Bars represent mean percent increase in contraction from unstimulated control with error bars indicating SEM (n = 7). Letters indicate differences as determined by a one way ANOVA and Newman-Keuls post hoc analysis (p < 0.05). D) Concentration-response curve for the inhibitory effects of carbenoxolone on crop contraction rates. Points represent percent inhibition at various log- 2 transformed concentrations with error bars representing SEM (n = 7, EC50 = 115.6 µM, R = 0.6613).

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3.2 Effects of serotonin, benzethonium chloride, and aedeskinin III on crop contraction rates

To shed light on the physiological regulation of crop contraction rates, we examined the effects of serotonin, benzethonium chloride, and aedeskinin III (AKIII).

Serotonin is an agonist of crop contractions in the blow fly, and is a general myoactive neurotransmitter in mosquitoes and other insects (Liscia et al. 2012). Benzethonium chloride is a mimic of drosomyosupressin, which inhibits crop contractions in the blowfly and suppresses muscle activity in other insects (Richer et al. 2000). AKIII stimulates hindgut contractions and the secretion of fluid by Malpighian tubules in mosquitoes

(Klaus W Beyenbach et al. 2009; Veenstra, Pattillo, and Petzel 1997). As shown in Fig. 27C, adding serotonin to the bath significantly stimulated crop contraction rates in a concentration dependent manner. A membrane permeable analog of cAMP (8-Bromo- cAMP; 1 mM) also significantly increased crop contraction rates, whereas the PKA inhibitor H-89 (10 µM) significantly decreases baseline contraction (p < 0.05, ANOVA with

Newman-Keuls post hoc analysis, n = 7) and prevents serotonin from increasing contraction (p > 0.05; ANOVA with Newman-Keuls post hoc analysis, n = 7). On the other hand, benzethonium chloride (25 µM) significantly elicited an 86.7 ± 9.7% inhibition of contraction rate (paired t-test, n = 7; p < 0.001). A full concentration-response curve for benzethonium chloride was not determined. The addition of AKIII to the bath did not

123 significantly change contraction rates at a concentration of 10 µM (paired t-test, n = 3; p

= 0.5601).

3.3 Pharmacological and molecular evidence for gap junctions in the crop

To determine the potential involvement of gap junctions in crop contractions, we examined the effects of CBX, which is a pharmacological inhibitor of gap junctions. As shown in Figure 27D, adding CBX to the bath inhibited crop contraction rates in a concentration dependent manner with an IC50 of 115.6 µM. Moreover, in crops that were pre-incubated with CBX (250 µM), the addition of serotonin (10 nM) did not significantly increase the contraction rate (n = 7; p > 0.05). Thus, the presence of active gap junctions appears to be critical to the baseline and serotonin-stimulated rates of crop contraction.

To confirm the presence of gap junctions, we quantified the molecular expression of innexin mRNAs in the crop. Innexins are genes that encode the gap junctional proteins of invertebrates; Ae. aegypti possesses six innexin (inx) genes: inx1, inx2, inx3, inx4, inx7, and inx8. As shown in Figure 28, inx2 was the most highly expressed innexin in the crop, followed by inx7. The expression of inx1, inx3, inx4, and inx8 mRNAs were also detectable, but at statistically lower levels than inx2 and inx7.

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Figure 28: Innexin mRNA relative expression levels in the Aedes aegypti crop. Values for each gene are expressed as relative expression to the reference gene, ribosomal protein S7 (RPS7). Bars represent the average relative expression ± SEM (n = 3). Letters indicate statistical significance as determined by one way ANOVA and Newman-Keuls post hoc analysis (P < 0.05).

In addition, we localized immunoreactivity for inx2 and inx3 in the crop of Ae. aegypti females using existing antibodies to Bombyx mori (Bm) inx2 and Aedes aegypti

(Ae) inx3. Notably, Bminx2-like immunoreactivity localized exclusively to the muscle cells of the crop as indicated by its colocalization with phalloidin (Figure 29A-C). The immunolabeling of inx2 occurred throughout the muscle fibers and was not restricted to intercellular junctions (Figure 29A). In contrast, Aeinx3 immunoreactivity localized 125 exclusively to the epithelial cells of the crop as indicated by its non-overlapping localization with phalloidin (Figure 29D-F). Notably, the labeling of inx3 was punctate or plaque-like near the lateral, intercellular membranes (Figure 29D), consistent with labeling of gap junctions (Bohrmann and Zimmermann 2008; Ostrowski, Bauer, and Hoch

2008).

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Figure 29: Inx2 (A-C) and inx3 (D-F) immunolocalization in the crop (ventral diverticulum) of adult female Aedes aegypti mosquitoes. Green indicates inx2 (A, C) or inx3 (D, F) immunolabeling. Red indicates actin labeling via phalloidin. Yellow in C and F indicates colocalization of inx and actin. Nuclei are counterstained with DAPI (cyan). Note that inx2 immunoreactivity primiarly coincides with phalloidin labeling of actin in the muscle cells, whereas inx3 labeling primarily occurs in the epithelium and does not overlap with actin.

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4. Discussion

The diverticulated crop is a storage organ conserved throughout insects of the order Diptera. The mechanisms by which the crop musculature contracts have been studied in the blow fly (P. regina), fruit fly (D. melanogaster), and house fly (M. domestica). Thus far, mosquito crops have been noted to contract and determined to be impermeable, but mechanisms of contraction have not been investigated (Clay et al.

1973; Nuttall and Shipley 1903). We utilized an in vitro assay to investigate the physiological regulation of crop contractions in mosquitoes. Our work demonstrates a conservation in the basic physiological mechanisms that regulate contractions in the crops of flies, and provides the first lines of evidence for a role of gap junctions in regulating contractions of the dipteran crop.

4.1 Mechanisms of crop contraction in mosquitoes and other dipterans

The present study has demonstrated that in vitro, the mosquito crop spontaneously contracts at a rate of ~5-15 contractions per min, and that these contractions require the presence of extracellular Ca2+. The baseline rate of contraction we measured in the mosquito is noticeably lower than those previously reported in P. regina, M. domestica, or D. melanogaster, where contraction rates range between 20 and

90 per minute, depending on crop volume (Haselton, Yin, and Stoffolano 2006; Kaminski et al. 2002; Liscia et al. 2012; Solari et al. 2013; Stoffolano, Danai, and Chambers 2013;

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Stoffolano, Patel, and Tran 2014). It is possible that the lower contraction rate in mosquitoes is related to their unique food handling compared to the other dipterans examined. That is, mosquitoes will only store sugar solutions in their crop, whereas blow flies store sugar and other essential nutrients (Day 1954; Stoffolano and Haselton 2013).

Similar to the crop of the blow fly (Liscia et al. 2012; Solari et al. 2013), extracellular

Ca2+ is essential for contractions of the mosquito crop. In the absence of extracellular Ca2+, the crop muscle cells are likely depleted of intracellular Ca2+, which would prevent the coupling of actin and mysion and thereby contraction (Szent-Györgyi 1975). Thus, calcium-dependence appears to be a conserved feature of crop muscle function in flies.

Data from our present study also indicate that contractions of the mosquito crop are under neuronal control. In particular, benzethonium chloride, a dromyosuppressin

(DMS) mimic (Lange et al. 1995), inhibits the contraction rates. Insect myosuppressins are neuropeptides that were first identified in the cockroach Leucophaea maderae, and shown to reduce hindgut contractions (Holman, Cook, and Nachman 1986). DMS, from D. melanogaster, was the first myosuppressin isolated and characterized in flies (Nichols

1991). The crop of the blow fly is innervated with DMS-immunoreactive neurons, and mRNAs encoding DMS receptors, DMSR-1 and DMSR-2, are highly-enriched in the crop of

D. melanogaster, with expression levels in the crop that are respectively over 100 and 10 times greater than in the whole animal (Chintapalli, Wang, and Dow 2007; Richer et al.

2000; Robinson et al. 2013). Moreover, DMS and benzethonium chloride both cause reductions of blow fly crop contraction rates in vitro (Richer et al. 2000). Thus, the

129 inhibitory effects of benzethonium chloride in the mosquito crop are likely due to activation of DMS receptors. It is unclear how DMS elicits its suppressive effect in the mosquito crop, but studies in D. melanogaster suggest that DMS acts via a GPCR that leads to a reduction of intracellular cAMP levels, presumably through coupling with an inhibitory G-protein (E. C. Johnson et al. 2003).

On the other hand, data from the present study indicate that contraction rates of the mosquito crop are stimulated by the neuronal monoamine, serotonin (5-HT).

Consistent with our physiological data, the crop of A. aegypti is innervated by serotonin- immunoreactive nerves, as well as FRMFamide- and cardioactive peptide b- immunoreactive nerves (Moffett and Moffett 2005). In D. melanogaster, the expression of mRNA encoding the serotonin receptor 5-HT7 is highly-enriched in the crop, at levels that are nearly 6-times greater than those in the while animal (Chintapalli et al., 2007;

Robinson et al., 2013). In A. aegypti, the orthologous receptor has been cloned from the

Malpighian tubules and demonstrated to signal via increasing intracellular cAMP (Lee and

Pietrantonio 2003; Pietrantonio, Jagge, and McDowell 2001). Consistent with this finding, we have shown that a membrane-permeant analog of cAMP also stimulates mosquito crop contraction rates.

Taking the above data together suggests that the crop contraction rates may be under ‘push-pull’ regulation via manipulation of intracellular concentrations of cAMP wherein DMS lowers cAMP levels and serotonin increases cAMP levels. Changes in cAMP levels may have down-stream effects on the activity of protein kinase A (PKA), which may

130 mediate the physiological effect via phosphorylation of proteins. Consistent with this notion, we have shown that inhibiting PKA with H-89 reduces baseline crop contraction rates and blocks the stimulation of crop contraction rates by serotonin. One potential downstream target of PKA may be gap junctions, which are known to be modulated by phosphorylation in systems as divergent as the vertebrate heart and fruit fly salivary glands (De Mello 1983, 1988). In fact, Stoffolano et al. (2010) suggested that the muscle cells of the blow fly crop may be electrically coupled, further suggesting a role of gap junctions in this muscle contraction.

4.2 Gap junctions and crop contraction:

Gap junctions are integral in contraction of the vertebrate heart and its pace making functions, and mediate electrical coupling in the Malpighian tubules of A. aegypti and salivary glands of D. melanogaster (Degroot et al. 2003; Hax, van Venrooij, and

Vossenberg 1974; Weng et al. 2008). As such, gap junctions could be playing similar roles in the contraction of the crop in mosquitoes. Supporting this hypothesis, the gap junction inhibitor carbenoxolone (CBX) inhibits mosquito crop contraction in a concentration- dependent fashion. The IC50 of CBX we determined (115.6 µM) is consistent with the concentrations of CBX commonly used to inhibit gap junctions composed of innexins in C. elegans and connexins in the mouse (Sangaletti, Dahl, and Bianchi 2014; Xia and Nawy

2003). Moreover, the addition of serotonin did not rescue the crop from the inhibition caused by CBX, suggesting that active intercellular communication/electrical coupling via

131 gap junctions is required for the both the spontaneous and serotonin-stimulated muscular contractions of the crop.

Consistent with our pharmacological results, two innexin mRNAs, inx2 and inx7, were abundantly expressed in the crop. Interestingly, the quantitative pattern of innexin gene expression in the crop most closely resembles that in the midgut and hindgut of adult females (Calkins et al., 2015, Calkins and Piermarini, in Review), suggesting that inx2 and inx7 are highly-expressed in tissues of alimentary canal. Notably, the immunoreactivity for inx2 localized exclusively to the network of muscle fibers in the crop, suggesting a specialized role in muscle function. The inx2 immunolabeling we find in the muscle cells of the crop, is strikingly similar to the labeling of inx11 in C. elegans, an innexin that is involved in electrical coupling of muscle cells in the C. elegans body wall (P. Liu et al.

2013). Moreover, inx2 has 4 predicted phosphorylation sites in the N-terminal region and an additional site in the C-terminal region of the predicted protein (Calkins et al. 2015); these sites could potentially be phosphorylated by kinases (e.g., PKA) to modulate gap junctional conductance. Thus, inx2 may be involved in the electrical coupling of muscle cells in the crop where it could play a role in coordinating the peristaltic contractions.

In contrast to inx2, inx3 discretely localizes to the intercellular boundaries between crop epithelial cells, suggesting that inx3 forms gap junctions and mediates intercellular communication between the epithelial cells. We have observed a similar localization pattern of inx3 in other epithelial tissues of adult female mosquitoes, including the hindgut, principal cells of the Malpighian tubules, and tracheal tubes of the

132 midgut (Calkins et al. 2015). Notably, the aforementioned tissues are all ectodermally derived, suggesting that inx3 may be particularly important in intercellular communication within ectodermally-derived tissues. Consistent with this notion, inx3 labeling is not detectable in the epithelium of the endodermally-derived midgut (Calkins et al. 2015). To date, the function of the crop epithelium is unknown. Presumably it is involved with secretion of cuticle that lines the lumen of the crop, but additional studies will be necessary to determine its functional significance and how Inx3 may contribute to intercellular communication within the epithelium.

5. Summary and Hypothetical model of crop muscle function

Integrating the data generated in this study with previous studies on gap junctions, we have developed a hypothetical model for the control of crop muscle contraction in mosquitoes (Figure 30). We propose a push-pull mechanism of cAMP signaling regulating crop contraction rates. In particular, we hypothesize that serotonin stimulates crop contraction rates by binding to a G-protein coupled receptor (GPCR) on muscle cells, potentially 5-HT7, leading to an activation of adenylyl cyclase and thereby an increase in intracellular cAMP. The increase of cAMP would activate protein kinase A

(PKA) to phosphorylate downstream proteins, which may include inx2. The phosphorylation of inx2 would open the gap junctions, thereby increasing cell electrical coupling and muscle contraction rates. On the other hand, we hypothesize that DMS

133 binds to an inhibitory G-coupled protein receptor, such as DMSR, causing an inhibition of adenylyl cyclase and cAMP production. The lowering of intracellular cAMP would lead to a decrease in PKA activity, thereby decreasing the phosphorylation of downstream proteins, such as inx2. Decreased phosphorylation of inx2 would likely lead to more gap junctions in the closed state, decreasing electrical coupling between cells and muscle contraction.

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Figure 30: Hypothetical model of crop muscle function based on work of present study. Schematic of the ventral diverticulum showing basic anatomy (left) with cross sectional view of the crop (right). Inx3 was localized to intercellular boundaries between epithelial cells, whereas Inx2 localized to the muscle cells of the crop. Our physiological studies suggest that 5-HT7 activates crop contractions, presumably via a G protein-coupled receptor (GPCR), that activates adenylyl cyclase, increasing intracellular cAMP levels, which would activate PKA that phosphorylates inx2, increasing gap junctional conductance between cells and increasing contractions. DMS on the other hand potentially activates an inhibitory GPCR that would inhibit adenylyl cyclase activity reducing intracellular cAMP levels, leading to decreased electrical coupling and reducing contractions.

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In summary, we demonstrate that contraction of the A. aegypti crop is under neuronal control similar to that of the blow fly, with crop contractions increased by serotonin and decreased by the DMS mimic, benzethonium chloride. Furthermore, as in the blow fly, extracellular Ca2+ is integral to crop contraction. We also demonstrate that

1) CBX, a gap junction inhibitor, impairs crop contraction, and 2) innexin-encoding genes are expressed in the crop, and 3) innexin immunoreactivity occurs in muscle and epithelial cells of the crop. Taken together, this work shows control mechanisms of contraction in the mosquito crop, illustrates a necessity of gap junctions in the crop and crop contraction, and provides a starting point for continued investigation into this historically understudied tissue of mosquitoes.

6. Acknowledgements

We thank Dr. Ryoichi Yoshimura of the Kyoto Institute of Technology, Department of Applied Biology, for his generous contribution of the inx2 antibody. We also thank Ms.

Nuris Acosta (OSU) and Ms. Edna Alfaro (OSU) for their assistance in mosquito rearing.

Funding for this study was provided by grants to 1) PMP from the NIH (R03DK090186) and

Mosquito Research Foundation (2014–03), and 2) TLC from the OARDC SEEDS program

(Grant# 2014–078; oardc.osu.edu/seeds), Ohio Mosquito Control Association Grant-In-

Aid, and Sigma Xi Grants in Aid of Research. State and Federal funds appropriated to the

OARDC of the Ohio State University also supported the study.

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Chapter 6: Synthesis

There are over 3,500 species of mosquitoes around the world, which are found on every continent except Antarctica (Harbach 2013). Only a few dozen of these species represent a threat to human or animal health, yet they are considered the most dangerous animals on Earth. The danger mosquitoes present is due to the diseases they transmit, and the burden of these diseases is severe; malaria alone causes over a half million yearly deaths, dengue afflicts hundreds of millions of people annually, and Zika is linked to serious birth defects when infecting pregnant women (Bhatt et al. 2013; Shao et al. 2016; Tang et al. 2016; WHO 2015). The mosquito subject of this study, Aedes aegypti, is the most important vector of the viruses that cause dengue, chikungunya, yellow, and

Zika fevers in humans. For many of these diseases, such as dengue and Zika, we do not currently have effective vaccinations, and instead must control mosquito populations to prevent spread of disease.

Mosquito control often relies on the use of insecticides targeting ion channels, synaptic enzymes, or synaptic receptors. Although these compounds are highly effective at killing mosquitoes, they exert a strong selective pressure driving evolution of resistance in the form of target-site and/or metabolic resistance. (Müller et al., 2008; Vontas et al.,

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2012). Thus, to ensure that our chemical arsenal for mosquito control remains viable, it is necessary to modify existing compounds to avoid resistance mechanisms, and/or identify new molecular and physiological insecticide targets with novel mechanisms of action.

Mosquito gap junctions could present one such target.

Gap junctions are intercellular channels that mediate direct communication via the transfer of certain small molecules and/or ions between adjacent cells. The gap junctions of vertebrates and invertebrates are composed of evolutionarily distinct families of protein subunits, known as connexins and innexins, respectively. Genomes of mosquitoes encode for six innexin proteins (Chapter 1). Despite their differences in protein composition, the gap junctions of vertebrates and invertebrates both play important roles in development (Giuliani et al. 2013; Iwamoto et al. 2013), the nervous system (Furshpan and Potter 1958; Revel and Karnovsky 1967; Revel, Olson, and

Karnovsky 1967), intercellular signaling (Bosco, Haefliger, and Meda 2011; Piermarini and

Calkins 2014), among others. Considering the importance of gap junctions in animals, and the evolutionary differences between vertebrate and invertebrate subunits; gap junctions may offer targets for vertebrate-safe insecticides, assuming the molecular differences could be exploited.

Pharmacological inhibition of gap junctions in adult female Aedes aegypti leads to incapacitation (flightless phenotype) and death of the mosquito, supporting the notion of targeting gap junctions for mosquito control (Chapter 2, Calkins and Piermarini, 2015).

Moreover, gap junction inhibitors kill larval mosquitoes, when added to their rearing

138 water (Chapter 2, Calkins and Piermarini, 2015). While this pharmacological inhibition used broad spectrum gap junction inhibitors, which block both vertebrate and invertebrate gap junction, these data proof the concept that gap junctions are potential targets for insecticide development. However, a strong basic understanding of physiological roles that innexins in mosquitoes could allow us to develop more specific insecticides as there are six innexin subunits that form potential mosquito gap junctions.

Gene expression and protein localization of innexins throughout the insect can act as a starting point for understanding the potential roles of innexins in mosquito physiology. Our studies of innexin gene expression in Aedes aegypti indicate that every tissue examined expresses at least 4 innexin genes suggesting the high molecular potential for homo- and heteromeric hemichannels, as well as homo- and heterotypic gap junctions (Chapter 3, 4, and 5; Calkins et al., 2015). Moreover, the expression of inx4 is highest in the gonads of both males and females (Chapter 3, Calkins et al., 2015), suggesting that inx4 may be playing similar roles in gonad development and gamete production in A. aegypti as in Anopheles gambiae and D. melanogaster (Tazuke et al.

2002; Thailayil et al. 2011). Additionally, inx8 is only detectable at the whole animal level in juvenile life stages (larvae and pupae; Chapter 3, Calkins et al., 2015), indicating that it may be playing similar roles in the mosquito’s postembryonic nervous system development as found in D. melanogaster (Curtin, Zhang, and Wyman 2002).

In addition to understanding baseline gene expression throughout the animal, expression changes after significant biological events may suggest functional importance.

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In adult female mosquitoes, consumption of a blood meal induces dramatic changes in gene expression in nearly every tissue that has been examined (Esquivel, Cassone, and

Piermarini 2014, 2016; Price et al. 2011; Sanders et al. 2013). Notably, we have shown that innexins are up-regulated 24-hours after a blood meal; in particular, inx2 is up- regulated in the whole insect (WI), Malpighian tubules (MT), Midgut (MG), and Ovaries

(OV), inx3 is up-regulated in the WI and OV, and inx4 is up-regulated in the WI (Chapter

4). Despite the upregulation of inx2 through several major tissues in the mosquito, knockdown resulted in no noticeable effect on survival or fecundity (Chapter 4). However, more research is necessary to determine if another phenotype is affected and/or to confirm that inx2 protein levels are reduced in the inx2-dsRNA injected mosquitoes. Thus, while the gene expression allows us to generate hypothesis on the potential physiological roles of gap junctions, understanding where the proteins localize to within a tissue offers an additional layer of resolution to potential innexin function.

For example, in D. melanogaster all eight innexins are expressed in the ovaries

(Robinson et al. 2013), however four innexins (inx1, inx2, inx3, and inx4) show discrete patterns of localization between the different cell types of the ovary (Bohrmann and

Zimmermann 2008). Similarly in A. aegypti inx3 localizes to intercellular boundaries between principal cells in the Malpighian tubules, and epithelial cells of both the crop and hindgut (Chapter 3, Chapter 5, Calkins et al., 2015). In the Malpighian tubules, the localization of inx3 between only principal cells and not at the membranes between stellate and principal cells (Chapter 3, Calkins et al., 2015), indicates that while inx3 is

140 likely involved in the coupling of principal cells, it is not likely to connect principal and stellate cells. Furthermore, in the crop, inx3 localizes to the epithelial cell layer, whereas inx2 localizes to the muscle cells, and while this doesn’t preclude an interaction between the two, it does suggest that these two innexins may be playing discrete roles in different tissue types within the crop (Chapter 5).

In addition to the molecular expression and immunolocalization of innexins that provide a starting point for hypotheses on functional innexin roles, in vivo and in vitro physiological studies, utilizing tools such as pharmacological agents, add further insights to potential physiological roles of gap junctions. In the Malpighian tubules of Aedes aegypti, principal cells are electrically coupled by gap junctions (Weng et al. 2008), with inx3 likely playing a role in this coupling (Chapter 3, Calkins et al., 2015). While the gap junctions between principal and stellate cells have not yet been identified, the kinin receptor, which triggers increased diuresis when activated, is located exclusively in the stellate cells (Lu, Kersch, and Pietrantonio 2011), despite stimulating principal cells (Klaus

W Beyenbach et al. 2009; Schepel et al. 2010). Furthermore, the majority of the metabolic

- activity that drives diuresis and produces HCO3 occurs in the principal cells while the

- HCO3 exchanger is located in the stellate cells (Piermarini et al. 2010). Taking the above information together strongly implicates gap junctions in both principal cell to principal cell connectivity, as well as principal cell to stellate cell connectivity, and an importance of gap junctions in diuresis. We have confirmed their importance in diuresis by

141 pharmacologically inhibiting gap junctions in vivo, which disrupts the diuretic capacity of adult females (Chapter 2, Calkins and Piermarini, 2015; Piermarini and Calkins, 2014).

Gap junctions are additionally physiologically important in the dipteran food storage organ, known as the ventral diverticulum or crop, which serves as a reservoir for the mosquito sugar meal (Day 1954; Stoffolano and Haselton 2013). Like other flies the mosquito crop goes through peristaltic contractions to pump food into the alimentary canal for digestion (Chapter 5, Thomson, 1975). The crop contraction is under neuronal control, with serotonin increasing contraction and dromyosuppresin decreasing contractions, presumably through the increase and decrease of cAMP, respectively

(Chapter 5, Johnson et al., 2003). Interestingly gap junctional conductance is known to be increased by increasing intracellular cAMP levels (Hax, van Venrooij, and Vossenberg

1974), and inhibition of gap junctions in the crop in vitro prevents contraction (Chapter

5). Taken into account with the localization of inx2 to the crop muscles, it seems likely that gap junctions (possibly comprised of inx2) are mediating electrical coupling integral to muscle contraction (Chapter 5). It would seem likely that this coupling is integral to other tissues that go through peristaltic contractions, such as the oviducts or hindgut, however this is yet to be determined.

Based on our current understanding of mosquito gap junctions it seems likely that these channels could provide good targets for the development of new insecticides. Gap junctions are likely playing roles in mosquito development, electrical synapses, reproduction, excretory physiology, and muscle contraction (Chapters 2-5, Calkins et al.

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2015; Calkins and Piermarini 2015; Piermarini and Calkins 2014). In the context of insecticides, one can envision developing inhibitors of gap junctions that arrest or disrupt development, thereby killing the insect. In the nervous system, abolishment of gap junctional communication at the electrical synapses could lead to loss of input to the CNS from receptors, such as those in the Johnston’s organ. In the reproductive system, chemical or genetic inhibition of innexins, inx4 particularly, could allow for a radiation free induction of sterility in males. Moreover, chemical inhibition in wild populations could impede egg production as well as sterilize adult mosquitoes. Inhibition of renal systems would lead to inability to reduce hemolymph osmotic stress after consumption of a blood meal and death of the mosquito. Finally, inhibition of muscle coupling, and thus muscle contraction would have serious system implications throughout the mosquito, potentially leading to paralysis and knockdown.

While gap junctions may make good insecticide targets, much work is still necessary before we reach that point. The electrical coupling of muscles in the crop is an excellent example of this. Stopping crop contraction would impede the mosquito’s ability to digest a sugar meal, reducing its longevity and fecundity (Foster 1995). Inhibiting contraction of muscles throughout the alimentary canal would prevent the movement of any food bolus through the mosquito’s gut, severely impeding longevity and fecundity.

But inhibiting muscle contraction throughout the whole insect would kill insects much the same way current neuronal-based insecticides do, however instead through direct action on the muscles. Thus it’s important to understand how widespread this coupling is in the

143 insect, to determine how effective of an insecticide we could develop. Moreover, the localization of inx3 to the intercellular boundaries in all ectodermally derived tissues is intriguing (Chapter 3 and Chapter 5, Calkins et al., 2015), but as of yet potential necessity of such extensive cell coupling is not well understood. As such, further investigation of inx3 function in these tissues is certainly warranted. Collectively, this body of work lays a framework for understanding the roles of gap junctions in the mosquito and demonstrates the potential for gap junctions as insecticidal targets.

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