the physiological roles of rhopr-kinins and the molecular ... · garima bhatt master of science...

The Physiological Roles of Rhopr-kinins and the Molecular Characterization of their Gene in the Blood-Gorging Insect,

Rhodnius prolixus

by

Garima Bhatt

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Cell and Systems Biology University of Toronto

© Copyright by Garima Bhatt 2012

ii

The Physiological Roles of Rhopr-kinins and the Molecular Characterization of their Gene

in the Blood-Gorging Insect, Rhodnius prolixus

Garima Bhatt

Master of Science

Department of Cell and Systems Biology University of Toronto

2012

Abstract

The dramatic feeding-related activities of the Chagas' disease vector, Rhodnius prolixus are

under neurohormonal regulation of serotonin and various neuropeptides. One such family of

neuropeptides, the insect kinins, possesses diuretic, digestive and myotropic activities in many

insects. In R. prolixus, they co-localize with the corticotropin-releasing factor (CRF)-like diuretic

hormone (DH) in neurosecretory cell bodies and their abdominal neurohaemal sites.

Additionally, kinins are present in endocrine cells of the midgut and are known to stimulate

hindgut and midgut contractions. Through the experimentation presented in this dissertation, the

cloning and spatial expression of the R. prolixus kinin (Rhopr-kinin) transcript is described.

Physiological bioassays demonstrate the myostimulatory effects of selected Rhopr-kinin peptides

and also illustrate the augmented responses of hindgut contractions to co-application of Rhopr-

kinin and Rhopr-CRF/DH. The irreversible effects of two synthetic kinin analogs on the hindgut

relative to the native kinins also exhibit the prospective biotechnological significance of this

study.

iii

Acknowledgments

We are very grateful to Dr. Ron Nachman for providing the kinin analogs and to the NSERC

Discovery Grant conferred to I.O., for supporting this research.

On a personal note, I would like to express my sincere thanks to my mentor and supervisor, Dr.

Ian Orchard, for granting me the wonderful opportunities to pursue research in his laboratory and

to interact with other experts in the field. His constant guidance as well as expertise in the field

of neuroendocrinology have made my research experience a thoroughly enriching one! Thank

you Dr. Orchard! I am also deeply grateful to Dr. Angela Lange and Dr. Tim Westwood for all

their help and advice during the entire course of my research.

All my Orchard and Lange lab mates, past and present, cannot be left out in this note of thanks!

Meet, DoHee, Lisa, Maryam, Laura, Marina, Jean-Paul, Vicki, Rose, and undergraduate

students: each one of you have made my graduate experience an exciting and rewarding one.

Thanks for all the support you have provided me during the roller coaster journey of my

research.

Lastly, special thanks to my parents, family and friends for their love, patience and for their trust

in my decisions despite the fact that it seemed like I was just isolating a gene or killing bugs!

iv

Table of Contents

Abstract………………………………………………………………………………….……….ii

Acknowledgements……………………………………………...………………………………iii

Organization of the Thesis………………………………………………………………..…….vi

List of Tables and Figures……………………………………………………………………...vii

List of Appended Tables and Figures………………………………..……………………….viii

List of Abbreviations…………………………………………………………………………....ix


in the Blood-Gorging Insect, Rhodnius prolixus……………………...……………...……..1

Introduction………………………………………………………………..………………….1

Rhodnius prolixus………………………………………………………………………...1

Salt and Water Balance………………………………………………………………….2

Communication through Neuroactive Chemicals………………………….……….….3

Neuropeptides and Other Factors…………………………….………………………...5

G Protein-Coupled Receptors (GPCRs)………………………………….…………….7

Neuropeptide Families Involved in Diuresis and Anti-diuresis in Insects….…..……8

CRF-related Peptides…………………………………………………....………………9

Insect Kinins…………………………………………………………………………….10

Objectives of Thesis…………………………………………………………………….18

v

Materials & Methods……………………………………………….……………………….19

Results……………………………………………………………....………………………..28

Expanded Discussion……………………………………………..………………….………53

Future Directions………………………………………………………….…………………64

References……………………………………………………………………………………66

Appendix: Molecular Identification and Characterization of Two Putative Serotonin Receptors in the Kissing Bug, Rhodnius prolixus………………………………..……..….81

Abstract………………………………………………………………………...…………….82

Introduction………………………………………………………………………….………83

Materials & Methods………………………………………………………………………..86

Results………………………………………………………………………………………..91

Discussion…………………………………………………………………….……………..113

Future Directions……………………………………………………...………………117

References…………………………………………………………………………………..118

vi

Organization of the Thesis

The body of this thesis is a comprehensive section of all the experimentation and findings of this

study on insect kinins in R. prolixus. This study will be submitted for publication shortly and

here, it includes expanded introduction and discussion sections.

The appendix includes a report on the experimentation and findings of a study done on serotonin

receptors in R. prolixus. I began my Masters working on these serotonin receptors but the

research was stalled and as a result, the topic was changed to the kinins. For completeness and

documentation, the preliminary results are shown here.

vii

List of Tables and Figures


in the Blood-Gorging Insect, Rhodnius prolixus……………………………………………..…1

Table 1. Summary of all the insect kinins that have been sequenced to date and their respective

physiological roles……………………………………………………...………………………..11

Table 2. Summary of the R. prolixus kinin (Rhopr-kinin) peptides and their associated peptides

previously sequenced through MALDI-TOF……………………………………….……………13

Table 3. List of important gene-specific primers used during the molecular cloning and

expression analysis of the Rhopr-kinin transcript…………………………………………….….21

Figure 1. A. Depiction of the nucleotide sequence and deduced open reading frame of the

Rhopr-kinin transcript. B. Structural composition of the Rhopr-kinin gene. C. Northern blot

analysis of R. prolixus central nervous system (CNS) RNA using a Rhopr-kinin probe….….…29

Figure 2. ClustalW2 alignment of Rhopr-kinin prepropeptide and other known and predicted

insect kinin prepropeptides……………………………………………………………………....31

Figure 3. Phylogenetic analysis of the Rhopr-kinin prepropeptide and other known and predicted

insect kinin prepropeptides…………………………………………………………………..…..33

Figure 4. Spatial expression profile of Rhopr-kinin transcript in 5th instar R. prolixus…..……..36

Figure 5. Localization of the Rhopr-kinin transcript to neurosecretory cells of the CNS of 5th

instar R. prolixus using a Rhopr-kinin probe…………………………………………………….38

Figure 6. Myotropic activities of 3 Rhopr-kinins on the 5th instar R. prolixus hindgut………....41

Figure 7. Synergism exhibited by co-application of a Rhopr-kinin and Rhopr-corticotropin-

releasing factor (CRF)-like diuretic hormone on the 5th instar R. prolixus hindgut……………..45

Figure 8. Effects of 2 synthetic kinin analogs on the hindgut of 5th instar R. prolixus….………50

Figure 9. Schematic displaying roles of kinins in the alimentary canal of R. prolixus……….…58

viii

List of Appended Tables and Figures

Table 1. List of gene-specific primers used for the molecular cloning and expression analysis of

the type 2 serotonin (5-HT2) receptor in R. prolixus based on the SK_Rhopr5HT2R Contig

generated by Song Kim…………………………………………………………………………..87

Table 2. List of gene-specific primers used for the molecular cloning and expression analysis of

a 5-HT2 receptor in the Malpighian tubules of 5th instar R. prolixus, with similarities to a protein

from Drosophila mojavensis…………………………………………………………..…………88

Figure 1. Depiction of the consensus sequence of the SK_Rhopr5HT2R Contig and its deduced

ORF………………………………………………………………………………..……………..92

Figure 2. Gel electrophoresis image of a PCR testing different primer combinations used to

clone cDNA segments that are part of the SK_Rhopr5HT2R Contig…………………...……….94

Figure 3. Spatial expression profile of the putative SK_Rhopr5HT2R transcript………...……..96

Figure 4. Post-prandial changes in expression of the putative SK_Rhopr5HT2R transcript…....98

Figure 5. Depiction of the consensus sequence of the GB_Rhopr5HT2R Contig and its deduced

ORF…..…………………………………………………………………………………...…….101

Figure 6. Gel electrophoresis image of a PCR testing different primer combinations used to

clone cDNA segments that are part of the GB_Rhopr5HT2R Contig………………….........….103

Figure 7. Spatial expression profile of the putative GB_Rhopr5HT2R transcript………….….107

Figure 8. A schematic displaying the genetic structure of the predicted gene encoded by the

GB_Rhopr5HT2R Contig……………………………………………………………………….109

Figure 9. ClustalW2 alignment of the putative GB_Rhopr5HT2R serotonin receptor and other

known and predicted insect serotonin receptors………………..………………………………111

ix

List of Abbreviations

Note: Only scientific words that are recurrently mentioned in this thesis are catalogued here. The abbreviations are stated following the first use of the word in the text. 5-HT: 5-hydroxytryptamine (also known as serotonin) ABN: abdominal nerve cAMP: 3’-5’-cyclic adenosine monophosphate CAP2b: cardioaccelatory peptide 2b CC: corpora cardiaca CNS: central nervous system CRF: corticotropin releasing factor DH: diuretic hormone DUM: dorsal unpaired medial (neurons) FISH: fluorescent in situ hybridization GPCR: G protein-coupled receptor MALDI-TOF MS/MS: matrix assisted laser desorption ionization time-of-flight tandem

mass spectrometry MTs: Malpighian tubules MTGM: mesothoracic ganglionic mass ORF: open reading frame PLNSC: posterior lateral neurosecretory cell PVO: perivisceral organ RACE: rapid amplification of cDNA ends Rhopr-kinin/K: Rhodnius prolixus kinin TEV: transepithelial voltage TMD: transmembrane domain

1


in the Blood-Gorging Insect, Rhodnius prolixus

Introduction

Rhodnius prolixus

The feeding-related physiology of the haematophagus hemipteran, Rhodnius prolixus has been

widely investigated ever since the pioneering work of the likes of insect physiologist, Sir Vincent

Wigglesworth, in the 1930s (Edwards, 1998). As a hemimetabolous insect belonging to the

Triatominae subfamily called “kissing bugs,” R. prolixus is an obligatory blood feeder, requiring

a blood meal to undergo moulting through each of 5 nymphal stages, followed by adulthood.

Interestingly, 5th instars can devour a large blood meal from their vertebrate hosts; a meal that

can weigh up to 10 times their original body weight. Carrying such a heavy load can be

evolutionarily disadvantageous for the species since it restricts mobility, making the insects

vulnerable to predation and being noticed by the host. Furthermore, the iso-osmotic haemolymph

of the insect is also compromised by the hypo-osmotic characteristics of the ingested blood meal.

Thus, immediately upon feeding (i.e. within 2-3 minutes), the kissing bug undergoes rapid

production and then elimination of urine, lasting 2-3 hours after feeding. These dramatic

physiological events make R. prolixus a fascinating organism to study in the field of insect

physiology. Concurrently, R. prolixus is 1 of 12 Triatominae species that play a medically

significant role in the transmission of the parasitic protozoan, Trypanosoma cruzi to the human

host (see Orchard, 2006). The proliferation of T. cruzi in the human body eventually develops

into full-fledged American trypanosomiasis or Chagas’ disease, affecting the nervous, cardiac

and gastroenterological systems (De Sousa, 2002). The insect is attracted to factors like odours,

temperature gradients, carbon dioxide and visual stimuli to reach a potential host (see Friend and

Smith, 1977). After consuming the blood meal from the human host, the insect may excrete the

parasite-harbouring urine onto the host. The victim may then rub the parasite into the eyes,

mouth and/or open wounds (WHO, 2008). First discovered by Brazilian Dr. Carlos Chagas in

1909, Chagas’ disease affects 15-18 million people globally and these statistics rise by 50,000

annually (see Lima et al.; 2010). This disease is considered an endemic and a neglected concern

2

in central and southern parts of the American continent. In fact, 1-6% of the Latin American

population is infected each year, with prevalence in destitute areas (see Dias et al.; 2002; Lima et

al.; 2010). Hence, acquiring more knowledge of the feeding-related events of R. prolixus is now

more important, as insight into its physiology and factors that regulate its systems may aid

pharmaceutical companies in their pursuit to mitigate the transmission of Chagas’ disease.

Salt and Water Balance

Organisms inhabiting warm and parched terrestrial environments are prone to desiccation.

Terrestrial insects possess small bodies with large surface area to volume ratios and may also

experience scarce food resources. Under these circumstances, the preservation of ionic and

osmotic homeostasis is fundamental for their survival. In fact, an adequate amount of water must

also be stored to facilitate development and reproduction (see Coast et al., 2002).

Osmoregulation is achieved when water gain and loss is balanced through physiological events

such as respiration, evaporation, metabolism, food consumption and excretion (see Coast et al.,

2002). Ingestion and excretion are directly linked since intake of excess amounts of water and

ions as well as metabolic wastes generated by the body must be excreted by the system. If this

were not the case, there would be disruptions in the composition and volume of the haemolymph,

which in turn would endanger the insect. Indeed, a 5th instar R. prolixus faces all of these

challenges along with the physical risk it encounters during the engorgement of a large blood

meal. When it is not experiencing post-prandial and possibly post-eclosion diuresis, R. prolixus

is under a natural state of anti-diuresis in order to conserve water, which is imperative since it

inhabits dry areas. During feeding, the hypo-osmolarity (~320 mosmol-1) of the blood meal risks

the iso-osmotic (~370 mosmol-1) balance of the insect haemolymph (Maddrell and Phillips,

1975). The immediate priority of the homeostatic system is to eliminate the plasma portion of the

blood meal which comprises excess amounts of water and ions through the process of rapid

diuresis. Once the blood meal enters the anterior midgut (crop), excess Na+ and Cl- ions are

absorbed into the haemolymph, while water follows via osmosis. As the nutritious component of

the blood meal concentrates in the midgut, the excess ions and water, now in the haemolymph,

are secreted into the distal Malpighian tubules (MTs). Here, the primary urine contains large

3

amounts of Na+ (125mmol-1) and potassium (K+) (70mmol-1) and yet is roughly iso-osmotic to

the haemolymph (see O’Donnell, 2009). As it makes its way to the proximal tubules, the urine is

further modified such that K+ and Cl- ions are selectively reabsorbed while water is not,

ultimately generating a hypo-osmotic urine (250 mosmol-1) that is passed into the hindgut and

eventually excreted out in frequent intervals, without further modifications (Maddrell, 1969;

Maddrell and Phillips, 1975). Meanwhile, the concentrated blood meal is in due course digested

as it passes down into the posterior midgut. Over several days after feeding, this disease vector

must conserve water and at the same time, eliminate excess ions (i.e. K+, Ca2+), organic anions,

and nitrogenous wastes that are generated from digesting the blood meal (see Coast, 2007;

O’Donnell, 2009). The entire process is a conglomeration of digestion, diuresis and anti-diuresis.

This leads us to question how these complex physiological events are controlled. The answer lies

with the synchronization of several neuroactive chemicals that serve as diuretic and anti-diuretic

hormones in the insect.

Communication through Neuroactive Chemicals

In the animal kingdom, cell to cell communication is a complex series of events that determines

the outcome of all multifaceted physiological, developmental and behavioural events. This

communication occurs through an ensemble of bioactive chemicals such as amino acids (e.g.

glutamate), lipid-based hormones (e.g. juvenile hormone), aminergic (e.g. serotonin) and

peptidergic compounds (e.g. insect kinins) that are transmitted between cells/organs (see Orchard

et al., 2001). Before the 1950s, it was generally accepted that specific endocrine cells or glands

release these chemicals. However, the discovery of neurosecretory cells by Ernst and Berta

Scharrer, in species like the teleost fish, Phoxinus laevis, and the woodroach, Leucophaea

maderae, revealed that neuronal activity was not simply electrical, rather, neurons can also

function as endocrine cells (see Scharrer, 1987; Guillemin, 2011). This discovery ultimately

opened doors to the field of neuroendocrinology.

Complex physiological events, such as ecdysis, reproduction, energy metabolism, digestion,

osmoregulation, etc; are biological phenomena that exhibit an integration of multiple biological

systems and exhibit accurate yet flexible outcomes. How is this accuracy and flexibility achieved

4

concurrently? The answer lies in the analysis of the neuroactive chemicals that drive such

cellular correspondence.

Bioactive chemicals originating from the nervous system, such as amines and neuropeptides, can

function in 3 ways: as (1) neurotransmitters (NTs), (2) neurohormones (NHs) and/or as (3)

neuromodulators (NMs); the same chemicals may also be released by endocrine cells at a

different location (e.g. midgut). As NTs, these neuroactive chemicals are released locally in a

synapse between the axon termini of a presynaptic neuron and a postsynaptic neuron or tissue

(e.g. muscle). The release of NTs is spatially exclusive, specific and is delivered quickly (see

Orchard et al., 2001; Nässel, 2009). Neurohormones on the other hand, are released into the

haemolymph from non-synaptic neurohaemal sites or varicosities located on peripheral nerves,

including abdominal nerves (ABNs), and forming, for example, perivisceral organs (PVOs) and

the corpora cardiaca (CC) (see Alstein and Nässel, 2010). Since insects have an open circulatory

system, the NHs are delivered to far reaches of the insect’s body. Hence, the response to NHs is

theoretically a slow, global and less-specific one. A neuromodulatory response is generally

somewhere between the NT and NH range and its action is generally on a larger region of tissues

(see Orchard et al., 2001; Nässel, 2002). Neuroactive factors can have 1 or all 3 of these roles,

although their function is solely dependent on whether ligand-sensitive receptors are present on

the target tissues.

Interestingly, these factors may be released individually or co-released with other factors to

instigate and coordinate intricate behaviours and also to alter them at appropriate times. These

chemicals could result in additive or synergistic effects on the target tissue. Thus, flexibility and

plasticity of the system is not only established by the variety of neuroactive chemicals but is also

determined by when, how and where they are released (see Orchard et al., 2001). Slight

differences in the structure of neuroactive chemicals (i.e. isoforms) may target different tissues

and if these are released during different biological events or time points in a specific event, the

system may allow for tweaking and adjustments to the overall phenotypic response.

5

Neuropeptides and Other Factors

Synthesis of Neuropeptides

The largest group of neuroactive chemicals in the animal kingdom is the neuropeptides.

Neuropeptides are encrypted in the genetic blueprint, even in biologically simple species such as

nematodes and Cnidarian species (see Hartenstein, 2006). The preprohormone or prepropeptide

transcript is translated in the rough endoplasmic reticulum and is then post-translationally

modified in the Golgi apparatus. Initially, the signal sequence is excised from the prohormone.

The active neuropeptides are generated when the prohormone is proteolytically cleaved at

specific mono-(Arg), di-(ArgLys, ArgArg, LysArg), and in rare cases tribasic (ArgArgArg,

LysArgArg) cleavage sites via prohormone convertases (PCs) (see Veenstra, 2000; Bendena,

2010). Since insect PCs and their respective cleavage sites are not well studied, most

contemporary studies employ knowledge of vertebrate cleavage sites to predict active peptides. It

is important to note that all predicted cleavage sites are not always functional, and so more

research on invertebrate PCs is required.

A majority of insect neuropeptides undergo α-amidation at the C-termini which is essentially

required for the functional activity of the neuropeptide during receptor binding. The precursors

for amidation have a Gly at the C-terminal followed by a cleavage site. Two enzymes synthesize

the amidated active peptide: peptidylglycine α-hydroxylating monooxygenase (PHM) and

peptidyl α-hydroxyglycine α-amidating lyase (PAL) (see Bendena 2010). The complexity of

peptide variation and regulation is enhanced by factors like gene duplication (i.e. paralogues),

alternative splicing and peptide modifications like disulfide bridge formation, sulfation, the

addition of a cyclic pyroglutamate at the N-terminus, etc; The bioactive neuropeptides are finally

packaged in 100-300nm, membrane-bound, electron-dense granules (see Nässel, 2002;

Hartenstein, 2006; Bendena, 2010).

Degradation of Neuropeptides

Several exopeptidases and endopeptidases have been identified in insects based on similar

activity found in vertebrates, such as the angiototensin converting enzyme (ACE), neprilysin

6

(NEP) and dipeptidyl-peptidase IV (DPPIV) (see Isaac et al., 2009). Susceptibility to inactivation

in the insect’s alimentary canal and the haemolymph can restrict the exploitation of

neuropeptides as pharmaceutical and research tools. Therefore, the use of ACE-inhibitors as well

as the development of ACE resistant synthetic analogs which contain factors like α-amino-

isobutyric acid (Aib), have been utilized as options in the past (Nachman et al., 2009; 2011).

Classification of Neuropeptides

The classification and nomenclature of neuropeptides and peptidergic hormones is a topic that

stirs debate. The conventional means of classification is generally based on homology of motifs

compared to other known vertebrate and invertebrate peptides, the peptide’s biological functions,

structure and localization (see Coast and Schooley, 2011). Nonetheless, we cannot rule out the

notion that a single peptide may have multiple species-specific functions and thus naming and

categorizing neuropeptides into families continues to be a complex task.

Experimental Approaches in Studying Neuropeptides

The first insect neuropeptide was isolated from the cockroach, Periplanata americana, in 1975

and was named proctolin, based on its ability to stimulate the hindgut (i.e. proctodeum). In 1988,

around 20 insect neuropeptides were known, but with the advent of state-of-the-art technology,

hundreds of insect neuropeptides have been identified (see Coast and Schooley, 2011). The

application of Matrix-Assisted Laser Desorption/Ionization Time of Flight (MALDI-TOF) mass

spectrometry, immunohistochemistry and various tissue specific biological assays (e.g. Ramsay

assay, myotropic assays, electrophysiology) have enabled neuroendocrinologists to not only

identify various neuropeptides, but also to study their expression in tissues and then to discover

their physiological relevance. Advancements in genomics and peptidomics have also fuelled the

discovery of neuropeptides and their transcripts from many insects (e.g. Predel and Neupert,

2008; Ons et al., 2009; Hauser et al., 2011). Furthermore, molecular techniques such as gene

cloning, Northern blot and in situ hybridizations, RNA interference, functional expression

assays, bioinformatics, etc; fuel our knowledge of how neuropeptides regulate physiological

7

events via cellular signal transduction. Also, the availability of invertebrate whole genome

assemblies, such as the recent assembly of the R. prolixus genomic database, has promoted

investigations on the role of neuropeptides and their receptors in this medically-relevant

hemipteran.

G Protein-Coupled Receptors (GPCRs)

Studies on receptors have been eminent in our understanding of how neuroactive chemicals

interact with cells. G protein-coupled receptors (GPCRs) are widely studied receptors in

vertebrates and invertebrates alike and are named based on their association to a variety of

heterotrimeric guanosine triphosphate (GTP)-binding proteins. GPCRs are ideal pharmaceutical

targets because they have a broad range of endogenous ligands like peptides, biogenic amines,

proteases, glycoproteins, nucleotides, ions and even photons (see Meeusen et al., 2003). The

superfamily of GPCRs comprises 7 α-helix transmembrane domains (TMDs) that are linked to

one another via 3 intracellular and 3 extracellular loops. The N-terminal is typically glycosylated

and is situated on the extracellular side whereas the C-terminal is located on the cytoplasmic end

and is involved in intracellular signal transduction. GPCRs are linked to numerous signal

transduction pathways involving phospholipase C, diacylglycerol as well as secondary

messengers like cyclic adenosine triphosphate (cAMP), inositol phosphates and Ca2+ ions (see

Meeusen et al., 2003; Claeys et al., 2005).

The availability of enormous computational and technological resources has enabled scientists

to pull out a large collection GPCRs based on their sequence homologies to GPCRs found in

other species. However, the lack of knowledge of their specific ligand places many GPCRs in a

category called “orphan receptors.” Various online 7 TMD prediction tools like the TMHMM

(http://www.cbs.dtu.dk/services/TMHMM-2.0/) on the ExPASy server further elucidate the

structural conformations of GPCRs and their sequence homologies (see Meeusen et al., 2003).

Moreover, in insects, GPCRs that specifically bind to neuroactive chemicals such as

neuropeptides and biogenic amines can be targeted pharmaceutically in order to control insect

disease vectors and agricultural pests (see Gäde and Goldsworthy, 2003). For this purpose,

artificially synthesized agonists and antagonists can also be employed to target specific GPCRs.

8

Neuropeptide Families Involved in Diuresis and Anti-diuresis in Insects

Diuretic Hormones

In insects, diuretic hormones target tissues like the midgut, the MTs, and the hindgut in order to

maintain ion and water balance. These diuretic hormones include the biogenic indole amine 5-

hydroxytryptamine (5-HT), insect kinins, corticotropin releasing factor (CRF)-related peptides,

the calcitonin (CT)-related peptides, and the cardioacceleratory peptide 2b (CAP2b)-related

peptides (see Coast, 2007; Coast and Garside, 2005).

The neurohormonal control of diuresis was first established by Simon Maddrell and colleagues in

the kissing bug, R. prolixus. The MTs in R. prolixus are not innervated and are stimulated to

secrete primary urine by haemolymph of fed insects (Maddrell, 1963). With this knowledge and

through skilful microdissections, Maddrell demonstrated that extracts of neurosecretory cells of

the mesothoracic ganglionic mass (MTGM) in the CNS stimulate the tubules and that the diuretic

hormone is released from neurohaemal sites on ABNs (Maddrell, 1964a; 1964b; 1966a; 1966b;

Berlind and Maddrell, 1979). One diuretic hormone was later identified as 5-HT (Maddrell et al.,

1991).

In R. prolixus, 5-HT is a diuretic hormone released into the haemolymph within 5 minutes of

feeding (Lange et al., 1989). Apart from increasing secretion (more than a 1000-fold) and

reabsorption by the MTs, 5-HT also has myotropic activity on the salivary glands, crop, dorsal

vessel and hindgut, while it also plasticizes the cuticle (see Orchard, 2006). 5-HT is co-localized

with the calcitonin-related DippuDH31 peptide (i.e. first isolated from Diploptera punctata) in the

dorsal unpaired medial neurons (DUM) of the MTGM and in the neurohaemal sites of all 5

ABNs (Te Brugge et al., 2005). DH31 increases the tubule secretion by a mere 17-fold (Te

Brugge et al., 2005). Diuresis in R. prolixus is stimulated by CRF-related peptides, which not

only display diuretic activity on tubule secretion, but also stimulate anterior midgut absorption

(Te Brugge et al., 2009). All of these diuretic factors stimulate hindgut and salivary gland

contraction in the kissing bug (see Orchard 2009). Interestingly, kinin-related peptides co-

localize in the posterior lateral neurosecretory cells (PLNSCs) of the MTGM with the CRF-

related peptides (Te Brugge et al., 2001). The co-localization of these diuretic hormones in the

respective neurosecretory cells demonstrates their potential to be released independently or

9

together according to the physiological requirements. The kinins possess diuretic activity in

several dipteran, lepidopteran and orthopteran species; however, in this hemipteran, they do not

stimulate tubule secretion, but do possess myotropic activity on the hindgut, salivary glands and

anterior midgut (see Torfs et al., 1999; Orchard, 2009; Coast, 2009).

Anti-diuretic Hormones

Fewer anti-diuretic hormones in insects are known in comparison to the exhaustive list of

diuretic hormones. Anti-diuretic factor (ADF) a and ADFb isolated from the mealworm beetle

Tenebrio molitor decrease fluid secretion of its MTs as well as that of Aedes aegypti (Eigenheer

et al., 2002; 2003; Massaro et al., 2004). In the locust, Schistocerca gregaria, the ion-transport

peptide (ITP) increases salt and water uptake from the ileum whereas the neuroparsins and

chloride transport stimulating hormone (CTSH) influence reabsorption from the rectum (see

Gäde, 2004). In R. prolixus however, the endogenous CAP2b-related peptide has a novel anti-

diuretic role as it decreases tubule secretion and also reduces salt and water absorption from the

crop (Paluzzi et al., 2008 and Ianowski et al., 2010).

CRF-related Peptides

First sequenced from the tobacco hornworm (Kataoka et al., 1989), the insect CRF-related

peptides are potent diuretic hormones (DH) that have been isolated from several species (e.g.

Kay et al., 1992; Baldwin et al., 2001). In the locust, the endogenous CRF-like peptide (i.e.

Locusta-DH) is immunolocalized in the neurohaemal sites of the CC and PVOs and increases

fluid secretion and cAMP levels of the MTs (Patel et al., 1994). In fact, there is also evidence for

its presence in the haemolymph, thereby making it a true diuretic neurohormone (Patel et al.,

1995; Audsley et al., 1997). These insect diuretic peptides are encompassed in a superfamily of

CRF peptides which are also prevalent in chordates, tunicates, molluscs and other phyla. The

vertebrate homologues of the insect CRF-like peptides include urotensin-1/urocortin/sauvagine,

urocortins 2 and 3 (Lovejoy and Barsyte-Lovejoy, 2010; see Lovejoy and Jahan, 2006).

10

In R. prolixus, the CRF-related peptide (i.e. Rhopr-CRF/DH) as well as the gene encoding it have

been sequenced by Te Brugge et al. (2011). Forty-nine amino acids in length, this peptide

stimulates the secretion rates of the MTs and also increases the absorption rates of the crop (Te

Brugge et al., 2011). Apart from being localized throughout the R. prolixus CNS, CRF-like

immunoreactivity was also found in nerve processes overlaying the posterior midgut and hindgut

(Te Brugge et al., 1999).

Insect Kinins

Insect kinins; not to be confused with the vertebrate kinins of the autacoids family, are a family

of neuropeptides found in arthropods, particularly insects. The kinins were first isolated from the

cockroach, Leucophaea maderae, hence were named leucokinins. The 8 isolated leucokinins

were then primarily known for their ability to stimulate hindgut contractions in L. maderae

(Holman et al., 1986ab; 1987ab). The family of insect kinins possesses a conserved C-terminal

pentapeptide motif: FX1X2WG-amide, whereby, X1 can be Ser, Phe, His, Asn, or Tyr and X2 can

be Ser, Pro or Ala. Over the years, endogenous kinins from several insect species have been

isolated and studied through a combination of high performance liquid chromatography (HPLC)

experiments, mass spectrometry and physiological bioassays (see Torfs et al., 1999).

Function of Kinins in Insects

The insect kinins have been identified in various insect species. Table 1 displays all the kinins

that have been identified and characterized so far, from insects, a crustacean and a molluscan

species. A majority of the insect kinins display potent myotropic activity on insect hindguts;

increasing amplitude and/or frequency of contractions. Although initially known for their

myotropic activity on the cockroach hindgut, the kinins are now also known for their diuretic

activity in several species. Studies have illustrated that kinins are able to increase fluid secretion

in most species studied, including the lepidopteran species, Manduca sexta (Blackburn et al.,

1995) and Heliothis virescens (Seinsche et al., 2000). In the dipteran species D. melanogaster

(O’Donnell et al., 1998) and A. aegypti (see Beyenbach and Piermarini, 2011) the kinins increase

11

Table 1. Summary of all sequenced insect kinins and their physiological activities. Conserved amino acids are bolded. Refer to Torfs et al., 1999 for detailed review. Functions listed below mainly focus on the myotropic and diuretic activity of peptides.

Phylum/ Class/Order

Species Peptide Name Amino acid Sequence Function Reference

Arthropoda / Insecta / Diptera

Aedes aegypti∞

Aedeskinin I NSKYVSKQKFYSWGa Myotropic activity on hindgut. Aedeskinins & CDPs depolarize TEV of MTs and increase secretion.

Veenstra et al., 1994; 1997; Schepel et al., 2010; Cady & Hagedorn, 1999ab; Chen et al., 1994∞

Aedeskinin II NPFHAYFSAWGa

Aedeskinin III NNPNVFYPWGa

Culex salinarius∞

CDP I NPFHSWGa Myotropic activity on L. maderae hindgut. Depolarize TEV of C. salinarius MTs.

Clottens et al., 1993∞; Hayes et al., 1994

CDP II NNANVFYPWGa CDP III TKYVSKQFFSWGa

Drosophila melanogaster∞ Drosokinin NSVVLGKKQRFHSWGa Increases fluid secretion of endogenous MTs.

Cantera et al., 1992∞; Terhzaz et al., 1999

Musca domestica∞ Muscakinin NTVVLGKKQRFHSWGa

Little myotropic activity on hindgut and none on midgut. Increases fluid secretion of endogenous MTs.

Iboni et al., 1998∞; Holman et al., 1999

Arthropoda / Insecta / Lepidoptera

Helicoverpa zea

Helicokinin I YFSPWGa Induce whole gut contractions. Possess in

vitro diuretic activity on M. sexta MTs by increasing fluid secretion but not in vivo.

Blackburn et al., 1995; Oeh & Nauen, 2003

Helicokinin II VRFSPWGa Helicokinin III KVKFSAWGa

Arthropoda / Insecta / Orthoptera

Acheta domesticus∞

Achetakinin I SGADFYPWGa Increase MTs fluid secretion and frequency of tubule movements. Possible synergism when applied with cAMP on MTs.

Coast et al., 1990; Chen et al., 1994a∞

Achetakinin II AYFSPWGa Achetakinin III ALPFSPWGa Achetakinin IV NFKFNPWGa Achetakinin V AFHSWGa

Locusta migratoria∞ Locustakinin AFSSWGa

Myotropic on L. maderae hindgut but not on endogenous hindgut or oviduct. Increases fluid secretion of L. migratoria MTs and rectum reabsorption. Also has diuretic activity on A. domesticus MTs.

Schoofs et al., 1992; Thompson et al., 1995∞

Arthropoda / Insecta / Blattaria

Leucophaea maderae∞

Leucokinin I DPAFNSWGa Potent myotropic activity on L. maderae hindgut but less effective on foregut and oviduct. No myotropic activity on the heart was present. Stimulates fluid secretion, depolarizes TEV and increase Cl- conductance in A. aegypti

MTs.

Holman et al., 1986ab; 1987ab; Nässel et al., 1992∞; Cook et al., 1989; 1990

Leucokinin II DPGFSSWGa Leucokinin III DQGFNSWGa Leucokinin IV DASFHSWGa Leucokinin V GSGFSSWGa Leucokinin VI pESSFHSWGa Leucokinin VII DPAFSSWGa Leucokinin VIII GADFYSWGa

12

∞ refers to immunohistochemical studies that have examined kinin-like reactivity in different species.

Periplanata Americana

Perikinin I RPSFASWGa

Stimulate hindgut contractions. Predel et al., 1997 Perikinin II DASFSSWGa Perikinin III DPSFNSWGa Perikinin IV GAQFSSWGa Perikinin V SPAFNSWGa

Arthropoda / Crustacea / Decapoda

Penaeus vannamei∞

Pev-kinin 1 ASFSPWGa

Myotropic on L. maderae and P. vannamei

hindguts. Increase fluid secretion in A. domesticus MTs.

Nieto et al., 1998∞; 1999

Pev-kinin 2 DFSAWAa Pev-kinin 3 PAFSPWGa Pev-kinin 4 VAFSPWGa Pev-kinin 5 pEAFSPWAa Pev-kinin 6 AFSPWAa

Mollusca / Gastropoda

Lymnaea stagnalis Lymnokinin PSFHSWSa

Physiological effects are not known. Cox et al., 1997

13

Table 2. Rhopr-kinins and kinin precursor/associated peptides that have been identified in CNS extracts via MALDI-TOF MS/MS by Te Brugge et al. (2011). Conserved amino acids are bolded. No. Peptide Name Peptide Sequence

1. RhoprK-1 TNNRGNFAGNPRMRFSSWAa 2. RhoprK-2 AKFSSWGa 3. RhoprK-3 ANKFSSWAa 4. RhoprK-4 AKFSSWAa 5. RhoprK-5 DEDRQKFSHWAa 6. RhoprK-6 GAKFSSWAa 7. RhoprK-7 AKFNSWGa 8. RhoprK-8 LSINPWKKIDDNGa 9. RhoprK-9 AKFSSWGa 10. RhoprK-10 ADDDWLKKARFNSWGa 11. RhoprK-109-15 ARFNSWGa 12. RhoprK-11 SAAAYTPLSWKRKPIFSSWGa 13. RhoprK-111-11 SAAAYTPLSW-OH 14. RhoprK-1113-20 KPIFSSWGa 15. RhoprK-1114-20 PIFSSWGa 16. RhoprK-12 RGPDFYAWGa 17. RhoprK-122-9 GPDFYAWGa 18. KPP-6 FSNEFMNDNNDIEKNIVEE-OH

14

the rate of primary urine production by augmenting the conductance for chloride towards the

tubule lumen. By increasing the secretion of primary urine, the insect kinins play a significant

role in ion and osmoregulation of these arthropods.

To date, studies on the function of insect kinins have been restricted to visceral muscle

contractions and diuresis. However, some research also sheds light on the roles of kinins in other

biological events. For instance, the kinins have been linked to the regulation of feeding

behaviour such as the meal size and frequency of feeding (Al-Anzi et al., 2010) and have also

been shown to be involved in olfactory and gustatory responses in D. melanogaster (Lopez-Arias

et al., 2011). Research on lepidopterans, H. zea (Nachman et al., 2003) and H. virescens

(Seinsche et al., 2000) illustrates the potential roles of kinins in osmoregulation and

development. Furthermore, in M. sexta, during the process of ecdysis, kinin along with DH30 and

DH41 work together as downstream peptidergic factors in orchestrating preecdysis I-like rhythm,

which is essential for shedding the old cuticle (Kim et al., 2006).

Immunolocalization of Insect Kinins

Leuckokinin I-like immunoreactivity has been detected in a wide range of insect species

representing Diptera, Lepidoptera, Orthoptera, Blattaria, etc; many of which are mentioned in

Table 1. Neuronal cell bodies have been detected throughout the CNS, including the brain,

subesophageal ganglion (SOG), thoracic ganglia and in the abdominal ganglia (see Torfs et al.,

1999). Leucokinin-like immunoreactivity has also been shown in nervous systems of the locust,

Schistocerca americana, the honey bee, Apis mellifera, the cockroach Nauphoeta cinerea (Chen

et al., 1994a), M. sexta (Chen et al., 1994b), in dipteran species Calliphora vomitoria, Phormia

terranovae (Cantera and Nässel, 1992) and Sarcophaga bullata (Sivasubramanian, 1994) as well

as in the lepidopteran species, Agrotis segetum (Cantera et al., 1992) and Spodoptera litura (Lee

et al., 1998). Species like the spider, Cupiennius salei (Schmid and Becherer, 1996) and the

nematode Ascaris suum (Smart et al., 1993) also have neurons that stain for leucokinins,

therefore suggesting that the insect kinins may have a greater evolutionary relevance than

previously thought (see Torfs et al., 1999). In L. maderae, kinin appears to act on the MTs and

15

the hindgut as a neurohormone because no leucokinin-like innervations have been seen. This is

supported by the presence of leukokinin-like neurohaemal sites and the lack of innervation to the

hindgut found in L. maderae suggesting that kinins function as neurohormones (Nässel et al.,

1992). Similarly, leucokinin stained-neurohaemal organs were also observed in A. domesticus

(Chen et al., 1994a).

The distribution of CRF-related peptides has also been studied in several of the above mentioned

species. Amongst them, M. sexta (Chen et al., 1994b) and L. migratoria (Thompson et al., 1995)

are two species in which kinin and CRF/DH are co-localized in the neurosecretory cells and

neurohaemal sites, therefore suggesting that these neuropeptides may be co-released and possibly

work in unison. This may be physiologically significant as in the case of the locust (Thompson et

al., 1995) and the housefly (Iboni et al., 1998), where there is evidence of synergism between

kinin and CRF on the tubule secretion rate.

Kinin Genes

Despite the advances in genomics, genes encoding the kinin preprohormones have only been

cloned from the dipteran species, A. aegypti (Veenstra et al., 1997), D. melanogaster (Terhzaz et

al., 1999) and from the moth, Bombyx mori (Roller et al., 2008). These species each possess a

single insect kinin gene, which varies in size. This can be attributed to the fact that the number of

kinin peptides encoded by these genes differs from species to species (Table 1). In silico searches

through genome and expressed sequence tag (EST) databases have confirmed the presence of the

kinin genes in Anopheles gambiae (Radford et al., 2004), the Monarch butterly, Danaus

plexippus (EHJ73323), in the aphid, Myzus persicae (Christie, 2008), 5 hymenopteran species

including Apis mellifera (Veenstra et al., 2012), in the water flea, Daphnia pulex (Gard et al.,

2009), the roundworm nematode, Trichinella spiralis (XP_003377687) and in a Tardigrada

species, Hypsibius dujardini (Christie et al., 2011). Bioinformatics analysis reveals that no kinin

precursor genes are present in the genomes of the red flour beetle Tribolium castaneum (Li et al.,

2008), nor of the parasitoid wasp, Nasonia vitripennis (Hauser et al., 2010).

16

Structural Attributes of Kinin

Structural analysis of the insect kinins reveals that the aromatic amino acids F1 and W4 are

essential in the functional activity of the peptide since these amino acids interact in a β-turn

conformation while interacting with the kinin receptor (Roberts et al., 1997). Furthermore, the

amidation at the C-terminal is also an important feature because a negatively charged amino acid

at the carboxyl terminus renders the peptide inactive (Nachman et al., 1995).

Kinin Receptor in Insects

A single kinin receptor has been cloned in A.aegypti (Pietrantonio et al., 2005), Anopheles

stephensi (Radford, et al; 2004), D. melanogaster (Radford et al., 2002), H. zea (Scherkenbeck,

et al; 2009), in the Southern cattle tick, Boophilus microplus (Holmes et al., 2000), the pond

snail L. stagnalis (Cox et al., 1997) and has also been predicted in the honey bee Apis mellifera

(Ac: XP_396025.3). Extensive analysis in A. aegypti indicates that the 3 endogenous kinins act

via a single multi-ligand receptor through different intracellular pathways (Pietrantonio et al.,

2005; Schepel et al., 2010). In the MTs of both, D. melanogaster and A. aegypti, the kinins and

their receptor activate the inositol triphosphate (IP3)-mediated Ca2+ pathway (Cady and

Hagedorn, 1999b; Radford et al., 2002; 2004; Pietrantonio et al., 2005). Additionally, the kinin

receptors have been immunologically localized to the stellate cells of the tubules in D.

melanogaster, A. stephensi and A. aegypti (Radford et al., 2002; 2004; Lu et al., 2011). Research

elucidates that kinins increase Cl- permeability through the stellate cells as well as through

septate junctions (Schepel et al., 2010; see Beyenbach and Piermarini, 2011). However, both the

paracellular and transcellular routes of anion flow are mediated by the stellate cells (Lu et al.,

2011).

In a recent study, the Aedes kinin receptor was immunolocalized in the head, midgut, tubules,

ovaries and hindgut (Kersch and Pietrantonio, 2011). The receptor was present in open-type

endocrine cells of the midgut, indicating again a potential digestive role of the kinins via a brain-

gut feedback system (Kersch and Pietrantonio, 2011; see Orchard et al.; 2001).

17

Kinins in Rhodnius prolixus: Rhopr-kinins

In R. prolixus, leucokinin I-like staining is found throughout the CNS (Te Brugge et al., 2001).

Leucokinin 1-like peptides co-localize with the CRF-like peptides in the posterior-lateral

neurosecretory cells (PLNSCs) of the MTGM, with staining also observed in ABNs 1 and 2 and

in endocrine cells of the midgut. Immunoreactivity for leucokinin-like peptides is also observed

in the proctodeal nerve in R. prolixus (Te Brugge et al., 2001). In the CNS, both CRF and kinin-

like immunoreactivity decreases in the PLNSCs and neurohaemal sites up to 2.5 hours after

feeding and these levels are restored 24 hours after feeding, further supporting the theory of co-

release (Te Brugge and Orchard, 2002b). In addition, leucokinin 1-like staining was also

observed in open-type endocrine cells of the midgut (i.e. possible paracrine role) and processes

were observed on the hindgut and posterior midgut (Te Brugge et al., 2001). Leucokinin 1 does

not appear to stimulate secretion by MTs, nor does it increase anterior midgut absorption (Te

Brugge et al., 2001; 2002ab; 2009). Nonetheless, leucokinin 1 depolarizes the midgut TEV and

resistance and also increases hindgut and midgut contractions, thereby suggesting its

involvement in digestion and excretion (Te Brugge et al., 2002ab; 2009). Interestingly, the R.

prolixus MTs are comprised of principal cells but lack stellate cells, and this may explain the

lack of response of the R. prolixus tubules to leucokinin. The endogenous kinin gene and its

receptor have not been isolated in R. prolixus. However, in silico studies have predicted the kinin

gene and have also confirmed the peptides encoded by it using MALDI-TOF MS/MS (Te

Brugge et al., 2010; Ons et al., 2011).

Table 2 displays the 18 predicted kinins and precursor associated (spacer) peptides encoded by

the Rhopr-kinin gene, and which were sequenced from the R. prolixus CNS by Te Brugge et.al.

(2010). The Rhopr-kinin gene encodes for the largest number of kinin peptides found thus far in

any species. Amongst these Rhopr-kinins are 5 atypical WA-amide kinins that have only

previously been found in a crustacean, P. vannamei (Nieto et al., 1998; 1999).

18

Objectives of Thesis

The objective of this thesis is to enhance our understanding of the physiological role of the

endogenous insect kinins in R. prolixus. The first aspect of this investigation was to isolate the

kinin gene transcript from the CNS and to map its expression patterns in 5th instar R. prolixus

through techniques in molecular biology. The physiological approach in this study was to

demonstrate a physiological function of 3 endogenous Rhopr-kinins and to examine for

synergistic effects of Rhopr-CRF/DH and Rhopr-kinin on contractions of the hindgut. Finally,

investigating the function of 2 kinin analogs on the R. prolixus hindgut added an additional

pharmacological facet to this project. The results of this study may be significant for

pharmaceutical industries pursuing an interest in the prevention of Chagas’ disease transmission,

by targeting digestion and/or excretion in this insect.

An appendix is also attached to elaborate on some research concerning the cloning of the

serotonin receptor in R. prolixus. This appended project was not completed successfully, but the

preliminary data is included here for future reference.

19

Materials & Methods

Animals

Rhodnius prolixus Stål were reared at 25°C at relatively high humidity in a well-established

colony at the University of Toronto Mississauga, ON, Canada. The insects were fed on

defibrinated rabbits’ blood (Cedarlane Laboratories Inc., Burlington, ON, Canada) at all post-

embryonic stages. Four to six week post-prandial, 5th instar R. prolixus of both sexes were used

for the experiments listed below. For molecular biology experiments, all tissues were dissected in

nuclease-free phosphate-buffered saline (PBS) (Sigma-Aldrich, Oakville, ON, Canada). Tissues

were immediately submerged in RNAlaterTM RNA stabilization reagent (Qiagen Inc.,

Mississauga, ON, Canada). For physiological assays, dissections were done in R. prolixus saline

modified from the protocol of Lane et al. (1975). The saline was composed of 150mM NaCl,

8.6mM KCl, 2.0 mM CaCl2, 4.0mM NaHCO3, 8.5 MgCl2, 0.02mM HEPES and 34mM glucose

in pH 7.0.

In Silico Search of R. prolixus Kinin Gene

A recent in silico study has predicted a cDNA encoding the Rhopr-kinin prepropeptide (Te

Brugge et al., 2010). Furthermore, Ons et al. (2010) have also identified a kinin prepropeptide in

R. prolixus. Based on these findings, the prepropeptide encoded by these cDNA sequences were

used to search the R. prolixus preliminary genome database via a tBLASTn analysis on Geneious

software 4.7.6 (Biomatters, Ltd., Aukland, New Zealand). Subsequently, the tBLASTn search

gave a positive hit in Contig 67 of the preliminary genome database. The putative Rhopr-kinin

gene’s promoter region (Reese, 2001) and potential splice sites (Reese et al., 1997) were then

predicted using respective online softwares by the Berkeley Drosophila Genome Project.

20

Isolation of the cDNA Encoding Rhopr-kinin Transcript

Based on the positive hits along the genomic database, multiple gene-specific primers were

designed and tested to amplify the Rhopr-kinin gene by screening a 5th instar CNS cDNA library.

Primers gbkininfor1 and gbkininrev1 (Table 3) were used to successfully amplify the entire open

reading frame (ORF) via Polymerase Chain Reaction (PCR) (Table 3). All PCR reactions were

performed using the S1000® Thermal Cycler (Bio-Rad Laboratories Ltd., Mississauga, ON,

Canada). The thermal cycler conditions for the PCR performed are listed sequentially: initial

denaturation at 95°C for 3 minutes, 30-35 cycles of denaturation at 94°C for 1 minute, annealing

at 58°C for 20 seconds, extension at 72°C for 30 seconds and final extension at 72°C for 10

minutes. PCR products were viewed through ethidium bromide stained 1% (w/v) agarose gels.

The PCR product of interest were either gel extracted via EZ-10 Spin Column DNA Gel

Extraction Kit (Bio Basic Inc., Markham, ON, Canada) or column purified using EZ-10 Spin

Column PCR Products Purification Kit (Bio Basic Inc., Markham, ON, Canada). The PCR

product was then cloned into the pGEM®-T vector (Promega Corporation, Madison, WI, USA)

after which, the recombinant vectors were transformed into competent Mach1™ T1R Escherichia

coli cells (Life Technologies Corporation, Carlsbad, CA, USA) that were grown into overnight

cell cultures. Plasmid DNA was then isolated from these cell cultures and sequenced at the

SickKids DNA Sequencing Facility (The Centre for Applied Genomics, Hospital for Sick

Children, Toronto, ON, Canada). A 5’/3’ Rapid Amplification of cDNA Ends (RACE) kit

(Roche Applied Science, Mannheim, Germany) was also employed in order to retrieve the 5’ and

3’ untranslated regions (UTRs).

Similar molecular cloning procedures were followed to clone PCR products that were used to

create an RNA probe for Northern blot and fluorescent in situ hybridization (FISH). The primers

used to amplify this product were kininfor1 and kininrev1 at an annealing temperature of 61°C

with similar thermal cycler conditions as mentioned above (Table 3). This product was also

cloned and sequenced to determine its directionality within the plasmid.

21

Table 3: Gene-specific primers used to amplify the Rhopr-kinin prepropeptide cDNA Oligo-nucleotide name Oligo-nucleotide sequence (5’-3’) gbkininfor1 AATGATTCTATTATGGATGGTATGG gbkininrev1 CGGTCAACGTAATGTAAATATTTAG kininfor1 GTAAACGTGATGAAGATAGGCAA kininrev1 GTGGTCCCTGAACAGATTGC Rhoprkininfor2 CACAGCTAAAGACA Rhoprkininrev3 GGCACCTCGTTTCTCTTCTTCA actinfor1 (positive control) ACACCCAGTTTTGCTTACGG actinrev1 (positive control) CACCTCGTTTCTCTTCTTCAAGC

Insect Kinin Phylogenetic Analysis

The sequenced Rhopr-kinin prepropeptide cDNA sequence was analyzed for a potential signal

peptide using the online SignalP 4.0 software (Petersen et al., 2011). A multiple sequence

alignment was generated of the R. prolixus kinin prepropeptide sequence (Ac: DAA34788) and

its homologs in other insect species using ClustalW2 (http://www.ebi.ac.uk/Tools/msa/clus-

talw2/). Prior to the alignment, online genomic and EST databases were searched using a

BLAST search with the Rhopr-kinin sequence as query on the ExPASy website

(http://ca.expasy.org/tools/dna.html) in order to identify potential homologs. Sequenced and

predicted insect kinin prepropeptide cDNAs were identified from the following species: B. mori

(Ac: BAG50367), D. plexippus (Ac: EHJ73323), A. aegypti (Ac: AAC47656), A. gambiae (in

silico prediction by Radford et al., 2004), C. quinquefasciatus (Ac: EDS35029), M. persicae (Ac:

ES224909), A. pisum (prediction based on M. persicae sequence), D. melanogaster (Ac:

NP_524893), D. ananassae (Ac: XP_001956413), D. erecta (Ac: XP_001972732), D. grimshawi

(Ac: XP_001984991), D. mojavensis (Ac: XP_002009025), D. persimilis (Ac: XP_002027015),

D. pseudoobscura (Ac: XP_001353697), D. sechellia (Ac: XP_002030488), D. simulans (Ac:

XP_002084857), D. virilis (Ac: XP_002047878), D. willistoni (Ac: XP_002074422) and D.

yakuba (Ac: XP_002094790). The multiple sequence alignment was exported to BOXSHADE

3.21 (http://www.ch.embnet.org/software/BOX_form.html) in order to generate a figure

highlighting amino acid conservations. The ClustalW2 alignment was then used to create an

22

unrooted phylogenetic tree via neighbour-joining (NJ) analysis with bootstrap values

appertaining to 1000 replicates. This analysis was done using MEGA5 (Tamura et al., 2011).

Assessment of Transcript Size via Northern Blot Hybridization

All materials used for RNA-related procedures were RNase-free. Total RNA was first extracted

from 100 CNS tissues via the Trizol® reagent (Life Technologies Corporation, Carlsbad, CA,

USA). 1µg of CNS total RNA was denatured at 75°C for 5 minutes and immediately chilled on

ice. Likewise, to determine the size of the transcript, 30ng of RiboRuler™ High Range RNA

Ladder (Fermentas Canada Inc., Burlington, ON, Canada) was also prepared. The RNA samples

and the RNA ladder was loaded with 2x RNA loading dye (Fermentas Canada Inc., Burlington,

ON, Canada) onto a 1% formaldehyde-agarose gel (2.2M formaldehyde, 20mM MOPS, 5mM

NaOAc, 1mM EDTA, pH 7.0). The RNA was electrophoresed for around 125 minutes at 70V.

To ensure the integrity of the RNA, the gel was viewed briefly under the UV transilluminator.

The gel was then rinsed in diethylpyrocarbonate (DEPC)-treated distilled water for 30 minutes to

remove any traces of formaldehyde which would otherwise result in high background. Through

an overnight process of downward capillary transfer, the RNA was transferred from the gel to a

positively charged nylon membrane (Roche, Mannheim, Germany) in 20X saline-sodium citrate

(SSC). The next day, the membrane was washed in DEPC-treated water for less than 10s. Next,

RNA was fixed onto the membrane via UV cross-linking at 30, 000µJ/cm2 (UVP CL-1000,

Upland, CA) and then dried at room temperature for 2-3 hours. Later, the membrane was stored

at 4°C. Prior to doing so, the gel was viewed under the UV to ensure successful RNA transfer to

the membrane. For the purposes of hybridization, the DIG-High Prime DNA Labeling and

Detection Starter Kit II (Roche, Mannheim, Germany) was used with some modifications made

to the manufacturer’s specifications. To specifically identify the Rhopr-kinin mRNA transcript

on the membrane, a digoxigenin (DIG)-labelled anti-sense RNA probe was created for

hybridization. 20µg of recombinant pGEM®-T vector containing a 563bp partial Rhopr-kinin

cDNA was linearized using the SacII restriction enzyme and was subsequently purified using the

EZ-10 Spin Column PCR Products Purification Kit (Bio Basic Inc., Markham, ON, Canada).

Next, the DIG RNA labelling kit SP6/T7 (Roche Applied Science, Mannheim, Germany) was

23

used to perform an in vitro transcription of 1µg of the SacII-linearized recombinant vector using

the SP6 RNA polymerase. The in vitro transcription reaction was then treated with

deoxyribonuclease I for 15 minutes at 37°C to remove the DNA template. The probe was stored

at -20°C for several weeks. A DNA DIG labelled probe was generated for the RNA ladder as per

manufacturers’ specifications for the labelling and detection kit. Prior to hybridization with the

probe, the nylon membrane was incubated with pre-warmed hybridization solution at 65°C for

45minutes in a process called prehybridization. Then, the prehybridization solution was replaced

with hybridization solution containing the Rhopr-kinin probe (~100ng/mL). The membrane was

incubated with the probe for 16-18hours at 65-68°C. Following hybridization, the membrane was

subjected to many stringency washes. First, the membrane was washed twice for 5 minutes in 2X

SSC, 0.1% SDS at 25°C followed by two washes for 15 minutes in 1X SSC, 0.1% SDS pre-

warmed to 65°C. During the immunological detection, Bioflex Scientific Imaging Films (Clonex

Corporation, Markham, ON, Canada) were exposed to the chemiluminescence substrate for

various times (1hour-24hours) at room temperature. To determine the size of the Rhopr-kinin

transcript, the RNA ladder was aligned with the Rhopr-kinin prepropeptide band. Three

independent biological replicates were performed.

Spatial Expression Mapping of Transcript via Reverse Transcription-Polymerase Chain

Reaction (RT-PCR)

Insects (equal number of males and females) were dissected and their tissues were pooled into 10

different groups: (1) CNS, (2) oesophagus, (3) salivary glands, (4) a pool of dorsal vessel,

trachea, fat body, (5) anterior midgut, (6) posterior midgut, (7) Malpighian tubules, (8) hindgut,

(9) ovaries and (10) testes. Total RNA was extracted using the Trizol® reagent (Life

Technologies Corporation, Carlsbad, CA, USA) and cDNA was then synthesized using

oligo(dT)20 primer that was provided by the iScript™ Select cDNA Synthesis kit (Bio-Rad

Laboratories Ltd., Mississauga, ON, Canada). The thermal cycler conditions for the cDNA

synthesis reaction are as follows: priming at 25°C for 5 minutes, reverse transcription at 42°C for

30 minutes and inactivation of enzyme at 85°C for 5 minutes. 400ng of RNA were used to

generate the cDNA, after which, it was diluted 1:2. An aliquot of cDNA was used as template in

24

a regular PCR to determine absence and presence of the Rhopr-kinin transcript in different

tissues. The Rhopr-kinin gene specific primers that were used in this experiment spanned over

the exon-exon boundary and were Rhoprkininfor2 and Rhoprkininrev3 (Table 3). For the

positive controls, actin primers actinfor1 and actinrev1 were used. The thermal cycler conditions

for this PCR were: initial denaturation at 95°C for 3 minutes, 28-30 cycles of denaturation at

94°C for 1 minute, annealing at 61°C for 20 seconds, extension at 72°C for 30 seconds and final

extension at 72°C for 10 minutes. This was repeated for 3 independent biological replicates.

Localization of the Rhopr-kinin Transcript in the CNS via Fluorescent In Situ Hybridization

(FISH)

The FISH protocol is specific for assessing cell-specific mRNA localization in R. prolixus and

was used as per previous studies (Paluzzi, et al; 2008) with some modifications. Additionally, for

the development of the FISH signal, Tyramide Signal Kit (TSA) #24 was used with some

modifications (Life Technologies Corporation, Carlsbad, CA, USA). The anti-sense DIG-

labelled Rhopr-kinin RNA probe generated for FISH is identical to the one used for the Northern

blot hybridization and was synthesized in the same way. For the negative control experiments, a

sense RNA probe was synthesized in the same manner but with T7 polymerase. Dissected

anterior and posterior midguts were submerged in 1X PBS and were then transferred into 4%

paraformaldehyde fixative for 30 minutes. The CNS tissues were left attached to the ventral

cuticle and directly submerged in the fixative for 30 minutes. The CNS tissues were then

dissected out into 1x PBS, 0.1% Tween 20 (PBST). All tissues were washed at least 5 times for 2

minutes each in PBST to remove all traces of fixative. To minimize background caused by the

endogenous peroxidase the tissues were quenched with 1% hydrogen peroxide at room

temperature for 10 minutes. Next, tissues were incubated with 4% Triton X-100 (Sigma-Aldrich,

Oakville, ON, Canada) in PBST for 1 hour at room temperature, to make the tissues permeable to

the RNA probes. The tissues were then washed several times with PBST to remove any

remaining Triton. The tissues were fixed again in 4% paraformaldehyde for 20 minutes. After

fixing, the tissues were once again, rinsed with PBST, to remove all traces of fixative. The

tissues were then rinsed in a 1:1 solution of PBST and RNA hybridization solution (50%

25

formamide, 5X SSC, 100µg/mL heparin, 100 µg/mL sonicated salmon sperm DNA and 0.1%

Tween 20) after which, they were transferred into 100% RNA hybridization solution.

Prehybridization solution was made by boiling RNA hybridization solution at 100°C for 5

minutes and then cooling on ice. Tissues were incubated with prehybridization solution at 56°C

for 2 hours. Meanwhile, 100ng of the RNA probe in 100µL of hybridization solution was

denatured at 80°C for 3 minutes and cooled on ice for 5 minutes. Tissues were incubated with the

RNA probe containing hybridization solution for 12-16 hours at 56°C. All wash solutions were

pre-warmed to 56°C and all washes were done at 56°C as well. The tissues were then washed

twice with pre-warmed hybridization solution for 15 minutes each, followed by3:1, 1:1 and 1:3

wash solutions composed of hybridization buffer: PBST respectively for 15 minutes each.

Finally, the tissues were washed 4 times with PBST for 5 minutes each. Next, the tissues were

blocked using 1x PBS, 0.1% Tween 20 and 1% blocking reagent (i.e. from TSA kit) (PBTB) for

10 minutes with constant mixing. A 1:400 dilution of the primary antibody, namely, Biotin-SP-

IgG Fraction Monoclonal Anti-Digoxin, was made in PBTB. The tissues were incubated with the

primary antibody for 2 hours are room temperature after which they were washed with PBTB 6

times for 10 minutes each. Next, tissues were incubated with the secondary antibody,

streptavidin-horseradish peroxidise in a 1:100 dilution (made in PBTB) for 1 hour at room

temperature. Tissues were washed 6 times with PBTB for 10 minutes each, once with PBST for

5 minutes and twice for 5 minutes each with 1X PBS, to remove any traces of the secondary

antibody. Finally, tissues were incubated with a 1:200 dilution of the Alexa Fluor® 568 tyramide

conjugate made in 1X PBS in the dark, at room temperature, for 2 hours. Tissues were then

rinsed several times with 1X PBS and washed in 1X PBS, overnight at 4°C. Tissues were

mounted the next day, in 100% glycerol and viewed under the epifluorescence microscope.

Hindgut Contraction Assays via Force Transducer

Dissections were done under R. prolixus physiological saline as mentioned earlier. All tissues

surrounding the hindgut (i.e. dorsal cuticle, dorsal vessel, diaphragm, MTs, trachea and fat) were

removed in order to expose the junction between the posterior midgut and the hindgut (i.e.

ampullae). Using a very fine silk thread, a double knot was tied at the junction of the posterior

26

midgut and hindgut. The posterior midgut above the knot was severed from the hindgut and the

hindgut was cut out of the insect along with a small portion of the ventral cuticle still attached to

the anus. The preparation was transferred to a well of a Sylgard-coated dish (Dow Corning,

Midland Michigan, USA; Paisley Products, Scarborough, ON) where the cuticle was pinned

down using minuten pins such that the hindgut was resting dorsal side up. The tissue was washed

with saline and then submerged in 200µL of saline. The loose end of the silk thread was then

attached (using a double knot) to the hook of a force transducer (Aksjeselskapet Mikro-

elektronikk, Horten, Norway) such that the hindgut was stretched in a physiologically

appropriate manner and the thread was at a 90° angle to the force transducer. The recordings

were translated onto a chart recorder (Linear 1200) via a strain gauge amplifier. The tissue was

initially equilibrated for 15 minutes. Next, the saline was replaced with either saline or different

synthetic peptides or analogs, based on the experiment. The amplitude of the peptide-induced

longitudinal contractions and the basal tonus were compared and measured. Three synthetic

Rhopr-kinins (GenScript, Piscataway, NJ, USA), were tested on the hindguts. The three Rhopr-

kinins were as follows: AKFSSWG-amide (K-2) at >95% purity, AKFNSWG-amide (K-7) at

>96.8% purity and AKFSSWA-amide (K-4) at >99.8% (Table 2). The Rhopr-kinin peptides

were stored as 10µL of 10-3M aliquots at -20°C. This was then used to make different doses

ranging from 5 x 10-6M to 1 x 10-11M via serial dilutions. Peptide doses were made and stored on

ice. Prior to application on the hindgut, each dose was warmed to room temperature and briefly

mixed using a vortex mixer. The doses were applied in a random order. After the response of

each dose was recorded, the preparation was washed with saline until the basal tonus returned to

control levels. At least 5 trials were performed for each dose tested for each Rhopr-kinin. To

analyze individual dose response curves as well as compare the responses of the 3 kinins at

individual doses, one-way analysis of variance (ANOVA) test was performed along with

Tukey’s multiple comparison tests (P<0.05).

In another series of hindgut contraction assays, two kinin analogs; 1728[Φ1]wp-3 ([Aib]FF

[Aib]WGa) and 1729 [Φ1]wp-1 (Ac-R[Aib]FF[Aib]WGa), were tested on the R. prolixus

hindgut at a concentration of 10-8M using the same methodology explained above. The kinin

analogs were supplied by Ron Nachman (USDA, College Station, TX, USA), in a collaborative

effort. Each kinin analog was tested separately on 5 independent trials and compared to Rhopr-

kinin 2. Analogs were stored at -4°C in 80% acetonitrile + 0.01% TFA, as 10-3M aliquots. Prior

27

to use, the analogs were dried and reconstituted in saline. Unlike the previous method, the

contractions were measured via Pico Oscilloscope 2202 and recorded via PicoLog (Pico

Technology, Cambridgeshire, UK). To determine the effectiveness of the kinin analogs in

comparison to their control kinin controls, one-tailed paired T-tests were done (P<0.05).

Hindgut Contraction Assays via Impedence

The hindgut was isolated as mentioned earlier. However, instead of tying the anterior end of the

hindgut with a thread, the hindgut was fixed onto a Sylgard-coated dish using minuten pins both

on the anterior end (i.e. at the ampullae) and the posterior end (i.e. on the ventral cuticle). The

hindgut was submerged in 200µL of saline. Electrodes were placed on either side of the anterior

region of the hindgut and peristaltic contractions were monitored via a chart recorder (Servogor

124) through a UFI impedance converter (Model 2991, Morro Bay, CA, USA). Concentrations

of 10-9M Rhopr-kinin 2 and 10-8M Rhopr-CRF/DH were applied individually, as well as together

on the hindgut, in order to deduce whether their effect is additive or synergistic. The doses were

prepared and applied in the same manner mentioned earlier. In each trace, each deflection was

counted as a contraction. To eliminate background noise, a line was drawn over the saline control

trace disregarding smaller deflections (i.e. which could also indicate spontaneous contractions).

This line was carried over the entire trace and only deflections surpassing the line were counted

as contractions. Both the amplitude and frequency of the contractions were measured in the

initial 10 seconds after application of the peptides and 6 independent trials were recorded. For

statistical analysis, one-way ANOVA was performed alongside Newman-Keuls multiple

comparison tests (P<0.05).

28

Results

Molecular Cloning and Sequencing of Rhopr-kinin cDNA

Using gene-specific primers based on the in silico prediction of the R. prolixus kinin gene (Ac:

BK007870; Te Brugge et al., 2010) and thereafter via 5’/3’ RACE, a 1251bp cDNA was

amplified (Figure 1A). The cloned cDNA is also identical to the Rhopr-kinin transcript predicted

by Ons et al., (2011; Ac: 1273279). The gene was determined to be structurally comprised of 2

exons that are 555bp and 642bp respectively and that are separated by an intron spanning 726bp

(Figure 1B). The online splice site prediction software as well as BLAST searches through the

Rhodnius genomic database have also confirmed these exon-intron boundaries (Reese et al.,

1997). The Northern blot hybridization using an anti-sense probe of 563bp in length consistently

yielded a single 2kb band (Figure 1C). Based on this prediction of transcript size, approximately

750bp of the 5’ and 3’ UTRs is still missing. Isolation of the UTRs has been unsuccessful thus

far; although a 36bp of the 5’ UTR and 18bp portion of the 3’ UTR have been sequenced. The

transcription start site and the polyadenylation signal (i.e. AATAAA) have yet to be isolated.

Nonetheless, an in silico prediction of the promoter region suggests that the transcription site is

approximately 652bp away from the translation start site (Reese et al., 2001). Multiple

polyadenlyation sites were also predicted in the 3’ UTR and the closest one is around 226bp

away from the translation stop site.

Rhopr-kinin Prepropeptide and its Phylogenetic Relevance

The sequenced Rhopr-kinin transcript encodes for a complete 398 amino acid prepropeptide

which consists of a 19 amino acid signal peptide that is cleaved between Gly19 and Asn20 in the

N-terminus (Petersen et al., 2011, Figure 1A). Twenty-four potential dibasic cleavage sites were

identified within the kinin prepropeptide; however, not all of these potential cleavage sites are

functional, as revealed by de novo sequencing of the kinin peptides via mass spectrometry

analysis of CNS extracts (Table 2; see Te Brugge et al., 2010). Fifteen identified kinin peptides

are amidated at the C-terminal and 2 precursor peptides are hydroxylated at the C-terminal

29

Figure 1. A is a depiction of the Rhopr-kinin cDNA and its single ORF encoding a complete

kinin prepropeptide sequence. The nucleotide base pairs (including the partial UTRs) and amino

acid sequences are numbered on the right. The start codon is bolded and italicized whereas the

stop codon is bolded and underlined. The sequence with the double underline represents the

signal peptide, with the cleavage site between Gly19 and Asn20. Dibasic cleavage sites are shaded

in grey and bolded regions are kinin peptides and kinin precursor peptides that are processed (Te

Brugge et al., 2010). C-terminal glycines required for amidation are boxed. The nucleotide

region highlighted in black represents the intron splice site. B illustrates a schematic of the kinin

gene comprising of 2 boxed exons and 1 intron between them. Both UTRs are represented as

lines on either ends of the exons. C displays the Northern blot hybridization using a 563bp anti-

sense kinin probe complementary to the coding sequence and was targeted against CNS RNA

(n=3 replicates). Size prediction as determined by molecular markers is indicated on the left.

31

Figure 2. A multiple sequence alignment displaying the comparison of Rhopr-kinin

prepropeptide to the kinin prepropeptides from other insects using ClustalW2. Kinin

prepropeptide sequences were retrieved from dipteran species D. melanogaster (Ac:

NP_524893), D. ananassae (Ac: XP_001956413), D. erecta (Ac: XP_001972732), D. grimshawi

(Ac: XP_001984991), D. mojavensis (Ac: XP_002009025), D. persimilis (Ac: XP_002027015),

D. pseudoobscura (Ac: XP_001353697), D. sechellia (Ac: XP_002030488), D. simulans (Ac:

XP_002084857), D. virilis (Ac: XP_002047878), D. willistoni (Ac: XP_002074422), D. yakuba

(Ac: XP_002094790), the mosquitoes, A. aegypti (Ac: AAC47656), A. gambiae (in silico

prediction by Radford et al., 2004), C. quinquefasciatus (Ac: EDS35029), the lepidopteran

species, B. mori (Ac: BAG50367), D. plexippus (Ac: EHJ73323) as well as the hemipterans, A.

pisum, and M. persicae (Ac: ES224909). Only amino acids that exhibit conservation in at least

50% of the column are highlighted: black represents identical amino acid and grey represents

similar conserved amino acids.

33

Figure 3. An unrooted phylogenetic tree displaying evolutionary relationships between the insect

kinin prepropeptide sequences from different orders, via neighbour-joining analysis with a

bootstrap value of a 1000. The branch lengths are drawn to scale and at each node, a bootstrap

percentage is provided. Bootstrap values less than 50% were not included here.

35

(Table 2). Amongst the predicted kinin peptides by Te Brugge et al., (2010), 10 peptides possess

a typical FX1X2WG-NH2 motif, 5 peptides have an atypical FX1X2WA-NH2 motif whereas the

precursor peptide K-8 has a DDNG-NH2 ending (Table 2; Figure 1A). The atypical WA-NH2

motif is so far unique to R. prolixus among the insects. Peptides K-10, K-11 and K-12 are

noteworthy because they contain internal dibasic cleavage sites which can also be cleaved to

generate smaller kinin peptides (Table 2).

The ClustalW2 multiple sequence alignment illustrates large variability within the kinin

prepropeptides of different insects. The sequences from Drosophila species have a high degree

of conservation. However, they are very different from the kinin prepropeptides from other

species, even mosquitoes. The kinin precursor sequences also vary in size, from Drosophila

species with a protein 160 amino acids in length, to the largest protein in R. prolixus which is

398 amino acids in length. Variability in prepropeptides is also created by the number of kinin

peptides they encode. For instance, the Drosophila prepropeptide encodes for 1 kinin, Aedes

possesses 3 kinins whereas the Rhodnius prepropeptide encodes for 16 kinin peptides (Table 1 &

2). The extent of the conservation is quite limited to the C-terminal motifs of the kinin peptides

(Figure 2).

Further phylogenetic analysis reveals that the kinin prepropeptides from the dipteran species

form a monophyletic group. Furthermore, the Rhodnius kinin prepropeptide forms a

monophyletic group with the kinin sequences of hemipterans and lepidopterans (Figure 3).

Spatial Expression Profile of the Rhopr-kinin Transcript

The qualitative spatial expression of the Rhopr-kinin transcript was determined via RT-PCR

using gene-specific primers that amplified a PCR product of approximately 580bp. Expression of

the kinin transcript was observed in the CNS of 4-6 weeks post-prandial 5th instars. Expression

was also observed in peripheral tissues such as hindgut, testes as well as in a pool of tissues

comprising of trachea, fat bodies and dorsal vessel (Figure 4). In some trials of RT-PCR,

expression of the kinin transcript was also observed in the midguts.

36

Figure 4. Spatial expression profile of Rhopr-kinin transcript. RNA was extracted from tissues

of 10, 6-week post-prandial 5th instar R. prolixus and cDNA was synthesized from each tissue

(n=3 replicates). A PCR was then run using gene-specific primers and the cDNA was used as

template. β-actin primers were used as a control. The results were viewed via gel electrophoresis.

Each lane represents a different cDNA template used: 1) CNS, 2) Salivary Glands, 3) Foregut, 4)

Anterior midgut, 5) Posterior midgut, 6) Dorsal vessel, Fat bodies & Trachea, 7) Malpighian

tubules, 8) Hindgut, 9) Ovaries, 10) Testes, 11) No template control.

38

Figure 5. Fluorescent in situ hybridization (FISH) of R. prolixus CNS using a 563bp anti-sense

kinin probe that is complementary to the kinin transcript. Hence, the Rhopr-kinin transcript was

localized to the PLNSCs of the mesothoracic ganglionic mass (MTGM) of the CNS. Preliminary

FISH was performed by Te Brugge and Orchard (unpublished).

40

Localization of Rhopr-kinin Transcript in CNS

Preliminary FISH results were performed using the same anti-sense probe utilized in the

Northern blot experiments. Previously, the kinin transcript was found to be expressed in neurons

within the CNS, especially in a cluster of bilaterally symmetrical posterior lateral neurosecretory

cells (PLNSCs) in the dorsal MTGM (Figure 5; Te Brugge and Orchard, unpublished). These

appear to be the same neurosecretory neurons that are positive for kinin-like immunoreactivity.

Currently, the FISH technique is being optimized to obtain a full expression map.

Physiological Effects of 3 Rhopr-kinins on Hindgut Contractions

All 3 Rhopr-kinins induced robust longitudinal muscle contractions, as revealed by the changes

in basal tonus (Figure 6A). The hindgut contraction assays testing these 3 Rhopr-kinins revealed

that all 3 peptides had myotropic activity on the hindgut muscle in a dose-dependent manner. At

lower concentrations ranging between 10-10M to a little over 10-8M, Rhopr-kinin 2 (K-2) appears

to be more effective at eliciting forceful contractions than Rhopr-kinin 7 (K-7) whereas Rhopr-

kinin 4 (K-4) was the least effective (Figures 6C). However, at concentrations above 10-8M, K-4

appears to induce the stronger contractions than the other 2 kinins. Analyses through one-way

ANOVA comparing doses for the individual kinins indicate that results are statistically

significant (Figure 6B). When the effects of the different peptides are compared at the same

doses via one-way ANOVA, the effects between the peptides are not statistically significant

(Figure 6C). It was also noticed that hindguts more full of digested material, responded more

robustly upon the application of these peptides in comparison to empty hindguts. The threshold

for K-2 is between 5x10-10M and 10-10M whereas the EC50 value is between 10-8M and 10-9M.

The threshold for both, K-4 and K-7 is between 10-9M and 10-10M. For K-4, the EC50 value is

around 10-8M whereas for K-7, it is a little higher than 10-8M (Figures 6C and D). Maximum

stimulation of the hindgut occurs at 10-6M for all 3 kinins although for K-2 and K-7, the graphs

indicate this value is 5x10-6M due to large values in single trials. The effects of all 3 kinins were

reversed when the tissue was washed in physiological saline.

41

Figure 6. A depicts example traces of hindgut contractions at different concentrations of kinins

tested using the force transducer. Upward arrows indicate the application of the peptides and the

downward arrows indicate the washing off of peptides. B individual dose response curves for

each Rhopr-kinin illustrating statistical significance based on one-way ANOVA paired with

Tukey’s multiple comparison tests (P<0.05). Each line represents doses that are not statistically

significance. Doses that are not encompassed within the same line or are not represented by a

line are statistically significant from one another. C is a dose response curve displaying the

changes in hindgut tension upon the application of different concentrations of 3 Rhopr-kinins. D

is a dose response curve displaying the effects of the 3 Rhopr-kinins through normalization as

percent of their own induced maximum hindgut basal tonus.

45

Figure 7. The co-operative effects of Rhopr-kinin 2 and Rhopr-CRF/DH on the hindgut of 5th

instar R. prolixus (n=5 replicates) tested via the impedence. A displays example traces of hindgut

contractions recorded via the impedence machine. B illustrates the changes in the amplitude of

contractions upon the addition of the peptides independently and then together. Likewise, C

illustrates the changes in frequency of contractions. One-way ANOVA and Newman-Keuls

multiple comparison tests (P<0.05) were done to determine statistical significance. Bars

annotated with the same letters represent no statistical significance.

48

Physiological Effects of Rhopr-kinin 2 and Rhopr-CRF/DH on Hindgut Contractions

Initially, the effects of Rhopr-kinin 2 and Rhopr-CRF/DH on the hindgut were tested on the force

transducer. However, it was visually detected that the peristaltic contractions produced by

Rhopr-CRF/DH were not picked up by the force transducer. Thus, the impedence machine was

used. Application of 10-8M Rhopr-CRF/DH produces robust contractions of the hindgut. It was

observed that the application of both peptides individually, increased the amplitude and

frequency of phasic contractions (Figure 7A). The effects of Rhopr-kinin 2 lasted briefly (i.e.

roughly 1 minute) whereas the contractions produced by the Rhopr-CRF/DH persisted for a

longer duration (i.e. around 3-5 minutes). The effects of Rhopr-CRF/DH were harder to wash off

the hindgut compared to the Rhopr-kinins, although these effects were still reversible (not

shown). Rhopr-CRF/DH appears to have a greater increase on the frequency of contractions

compared to Rhopr-kinin 2, although the results are not statistically significant (Figure 7C).

The application of 10-9M Rhopr-kinin 2 and 10-8M Rhopr-CRF/DH together on the hindgut

resulted in robust contractions with increased amplitude and frequency of the contractions

compared to the peptides applied alone (Figure 7B and C). In some trials, it was noted that the

frequency of contractions decreased but the amplitude of the contractions were bigger and the

contractions were more robust (Figure 7A).

Physiological Effects of the Kinin Analogs

Both Aib-containing kinin analogs at 10-8M increased basal tonus of the hindgut in a similar

manner to the Rhopr-kinins although both analogs produced significantly larger contractions in

comparison to Rhopr-kinin 2 (Figure 8A and B). Kinin analog 1729[Φ1]wp-1 had a larger effect

on the amplitude of the contractions than analog 1728[Φ1]wp-3.

Unlike the endogenous peptides, both kinin analogs were very difficult to wash off the hindgut.

In fact, the effects of the analogs were relatively irreversible and the basal tonus could not be

49

returned to the baseline in the time course used (Figure 8Ai and Bi). Hence, the effects of both

the kinin analogs could not be compared on the same hindgut preparation.

50

Figure 8. The effects of 2 Aib-containing kinin analogs at a concentration of 10-8M on the

hindgut, in comparison to Rhopr-kinin 2 (n = 5 replicates/analog), tested on the force transducer.

A illustrates the myostimulatory effects of the kinin analog 1728[Φ1]wp-3 whereas B displays

the effects of 1729[Φ1]wp-1 and each of them is compared to its respective control Rhopr-kinin

2. For both kinin analogs, i show example traces where upward arrows indicate the application of

the peptides and the downward arrows indicate the removal of peptide from the hindgut

preparation using saline washes. ii illustrates bar graphs analyzing the changes in the effects of

the analogs compared to the endogenous kinin, on the tension of the hindgut. Statistical analysis

was performed using one-tailed paired T-tests (P<0.05).

53

Discussion

The enterprising fields of genomics and peptidomics are constantly supplying information of the

presence and absence of neuropeptides and their genes in different species. However, little is

known about the species-specific functions of numerous neuropeptides. The molecular cloning

and expression mapping of the insect kinin cDNA from R. prolixus in this study, is a first in any

hemipteran species. The cloned insect kinin cDNA is identical to the in silico prediction (Ac:

BK007870) by Te Brugge et al., (2010) and encodes for the full ORF. Based on the Northern

blot prediction, the Rhopr-kinin transcript is approximately 2kb in length and thus, the Rhopr-

kinin cDNA is not complete and is missing around 750bp of the 5’ and 3’ UTRs (Figure 1).

Screening the 5th instar CNS cDNA library and 5’ and 3’ RACE were both unsuccessful in

yielding the complete UTRs and this is most likely due to poor primer designs. Thus, 5’ and 3’

RACE need to be repeated in the future, using more efficient primers. Although multiple

polyadenylation signals have been predicted in the 3’ UTR, all except the closest signal can be

ruled out since only a single band was observed in the Northern blot hybridization (Figure 1C).

This observation also rules out any other possibility of alternative splicing. Furthermore, BLAST

searches along the genomic database also dismiss any possibility of gene duplications because a

single hit region was obtained. Nevertheless, a Southern blot could be performed in the future to

confirm the presence of a single kinin gene in R. prolixus. Thus far, a single insect kinin gene has

been isolated from A. aegypti (Veenstra et al., 1997), D. melanogaster (Terhzaz et al., 1999) and

B. mori (Roller et al., 2008) and has been predicted from many more organisms (Radford et al.,

2004; Christie, 2008; Gard et al., 2009). The kinin gene from these insects also produces a single

transcript variant and hence these preliminary studies imply that evolutionarily, no gene

duplication or alternative splicing events have occurred within the insect kinin genes. The kinin

gene consists of 2 exons (Figure 1B) although little is known about the kinin gene structures in

other species.

Amongst the kinin precursor genes isolated and predicted, the Rhopr-kinin gene is the largest.

The differences in the sizes of the kinin genes can be accounted for by the number of kinin

peptide isoforms found in each species (Table 1). For instance, the kinin gene of D.

melanogaster encodes for a single kinin peptide (Terhzaz et al., 1999), the gene in mosquitoes

54

like A. aegypti (Veenstra et al., 1997) and A. gambiae (Radford et al., 2004) encodes for 3

peptides whereas in R. prolixus, 15 kinin peptides are encoded by the Rhopr-kinin gene (Table

2). The variability in size and the differences in the number of kinin isoforms encoded by these

prepropeptides explain the lack of conservation observed in the multiple sequence alignment

(Figure 2). Phylogenetic analyses reveal that the Rhopr-kinin sequence shares similarities with a

monophyletic group comprising hemipteran and lepidopteran kinin prepropeptides. On the other

hand, the dipteran kinin sequences form a monophyletic group thereby illustrating the

evolutionary conservation of the kinin sequences amongst the fruit fly species and the mosquito

species respectively (Figure 3). It is interesting to note that the transcripts encoding insect kinin-

like peptides have been predicted in other ecdysozoan species such as in the nematode, T.

spiralis and in a Tardigrada species, H. dujardini (Christie et al., 2011) whereas none were found

in the genomes of T. castaneum (Li et al., 2008) and N. vitripennis (Hauser et al., 2010). This

alludes to the fact that a predecessor to the insect kinins may have originated in the earlier stages

of ecdysozoan evolution and over time, certain insects may have possibly lost the kinin gene, and

in so doing, presumably developed alternative strategies in diuresis and digestion. Further insight

into the origins of the insect kinins can only be elucidated through further genomic studies as

well as by studying their physiological roles in different species.

Through mass spectrometry analysis, numerous Rhopr-kinins that are post-translationally

processed from the prepropeptide have already been identified (Te Brugge 2010; Table 2).

Amongst these Rhopr-kinin peptides are 6 peptides that have the typical FX1X2WG-NH2 C-

terminal pentapeptide motif. Amongst them are 3 Rhopr-kinins that may be further processed

into smaller peptides, making a total of 10 typical kinin peptides. Intriguingly, the prepropeptide

also processes 5 atypical kinins with the C-terminal pentapeptide motif, FX1X2WA-NH2 (Table

2). This is unique within the insect kinins and has only been found once before in a crustacean

white shrimp, P. vannamei (Nieto et al., 1998; 1999). Three kinin associated (spacer) peptides

that do not possess the C-terminal pentapeptide motif were also identified. Two of these peptides

(K-111-11 and KPP-6) are hydroxylated at the C-terminal whereas K-8 has a DDNG-NH2 ending

(Table 2). None of the other kinin precursor peptides predicted in the prepropeptide sequence

were identified (Te Brugge et al., 2010; Figure 1A). More research into prohormone convertases

is required to understand why certain dibasic cleavage sites may or may not be excised.

55

The spatial expression profile of the Rhopr-kinin transcript reveals that it is expressed in the CNS

of 4-6 weeks post-prandial 5th instars (Figure 4). This matches the preliminary FISH results

which reveal that the Rhopr-kinin transcript is present in the neurosecretory cells (PLNSCs) of

the dorsal MTGM (Figure 5) and this is consistent with the immunoreactivity observed in these

same cells (Te Brugge et al., 2001). The in situ hybridization results validate the synthesis and

production of the Rhopr-kinins by the PLNSCs. More trials of FISH need to be repeated to locate

other neurons that produce the Rhopr-kinin transcript in the CNS. The insect kinins have been

localized to the CNS of numerous insects via immunohistochemistry (see Introduction; Table 1).

Expression in the midgut using RT-PCR also matches with the leucokinin I-like

immunoreactivity found in open-type endocrine cells in the posterior regions of the midgut (Te

Brugge et al., 2001). Positive staining for kinin-like immunoreactivity in the midgut is limited to

R. prolixus although the kinin receptor has been localized to the posterior regions of the A.

aegypti midgut (Kersch and Pietrantonio, 2011). The localization of the Rhopr-kinin peptides to

the endocrine cells of the midgut indicates that the kinins may be involved in meal digestion (e.g.

by initiating digestive enzyme release into gut) and may also be associated with a brain-gut

feedback system (see Orchard et al., 2001). Relative to the CNS, lower levels of expression of

the Rhopr-kinin transcript is observed in the hindgut, the testes and a pool of dorsal vessel,

trachea and fat body (Figure 4). No kinin-like immunoreactive endocrine cells have been

previously discovered in these tissues, although leucokinin I-like immunoreactive processes have

been observed over the hindgut and posterior midgut (Te Brugge et al., 2001). It is highly

plausible that peripheral neurons that contain Rhopr-kinins may be associated with these tissues

and were as result detected in the RT-PCR. Several other neuropeptides are known to be

expressed in peripheral neurons (e.g. Gonzalez and Orchard, 2008; Te Brugge et al., 1999; Tsang

and Orchard, 1991). Likewise, the expression in the testes could also be due to Rhopr-kinin

containing-neurons associated with the testes. This corroborates with the findings that

leucokinin-1 staining has been observed in the genital nerve of R. prolixus (Te Brugge et al.,

2001). However, only further in situ hybridization and immunohistochemistry studies on these

tissues can verify the location of the kinin transcript and peptides. The presence of the Rhopr-

kinin transcript in the pool of trachea, fat body and dorsal vessel could be justified with the

possibility of peripheral endocrine cells like the Inka cells of the epitracheal glands in M. sexta

56

(Žitňan et al., 1996) that may produce the Rhopr-kinins. Again, further investigation is required

here.

Considering the leucokinin I-like innervation over the R. prolixus hindgut, 3 endogenous kinins,

Rhopr-kinins 2, 4 and 7 were synthesized and tested to understand their physiological role on the

hindgut. All 3 Rhopr-kinins effectively increase longitudinal muscle contractions of the hindgut.

Furthermore, it appears that Rhopr-kinin 2 is more effective than Rhopr-kinin 7 and the latter is

more effective than Rhopr-kinin 4 at concentrations between 10-10M and 10-8M (Figure 6C). At

concentrations above 10-8M, Rhopr-kinin 4 appears to elicit stronger contractions than the other

2 native kinins. Although these differences between the kinins are not statistically significant,

they can most likely be attributed to the differences in their amino acid sequences which perhaps

slightly alter peptide-receptor binding and thus potentially, receptor signaling. Rhopr-kinin 2 is

AKFSSWG-NH2 whereas in Rhopr-kinin 7, the Ser of the 4th residue is replaced by Asn. The

atypical Rhopr-kinin 4 is identical to Rhopr-kinin 2, except, the Gly of the 7th residue is replaced

by Ala (Table 2). The unusual Rhopr-kinin 4 is similar to the kinins identified in the crustacean

shrimp, Penaeus vannamei. The endogenous shrimp kinins are biologically active in stimulating

the cockroach hindgut contractions as well as in stimulating MTs secretions of the cricket but are

not as effective as the typical kinins (Nieto et al., 1998). This is in agreement with our findings.

Nonetheless, it is important to note that there is no statistical difference in the activities of these

peptides at the same doses, therefore these selected Rhopr-kinins are most likely functionally

redundant. However, there are multiple other Rhopr-kinins, of varied lengths and composition

that have yet to be tested. Experimentation of all the endogenous kinins will enable us to

determine whether there are differences in peptide potencies in their myotropic activity on the

hindgut.

The robustness of the Rhopr-kinin-induced contractions of the hindgut also increased when the

hindgut was fuller of digested material. It is possible that the muscle is stretch sensitive and thus

enhances its sensitivity to the Rhopr-kinins; thereby, producing bigger and more robust

contractions. These Rhopr-kinins also produce similar myostimulatory effects on the anterior

midgut (Orchard and da Silva, unpublished). A previous study has shown that leucokinin I does

not stimulate anterior midgut absorption although it does depolarize the TEV and alter electrical

resistance of the midgut (Te Brugge et al., 2009). The myotropic activity of the Rhopr-kinins on

57

the hindgut and the midgut not only demonstrate their potential roles in digestion and excretion,

but also exemplify their importance in overall feeding-associated activities. For instance, robust

and rhythmic contractions of these large tissues can function as accessory hearts, which will

effectively circulate neurohormones and nutrients throughout the insect system while also mixing

the contents of the crop.

58

Figure 9. Diagram displaying the roles of native and non-native kinins in the alimentary canal of

R. prolixus (present study; Te Brugge et al., 2001; 2002ab; 2009; Orchard and Te Brugge, 2002;

see Orchard, 2006; 2009). Modifications made to illustration by Zach McLaughlin.

60

A model (figure 9) depicts the known functions of insect kinins on the alimentary system of 5th

instar R. prolixus. Effects of insect kinins on the foregut are unknown. Leucokinin I does

stimulate contractions of principal salivary gland muscles but no kinin-like innervation is

observed in nerve processes overlaying the glands, suggesting a possible neurohormonal role

(Orchard and Te Brugge, 2002). The secretions of the MTs are not affected by leucokinin I (Te

Brugge et al., 2001; 2002ab; 2009) which is unique to this insect considering their diuretic

function in several insects (see Introduction; Table 1). In dipertan species like A. aegypti (Lu et

al., 2011), A. stephensi (Radford et al., 2004) and D. melanogaster (Radford et al., 2002), the

kinin receptor has been localized to the stellate cells of the tubules where they are involved in Cl-

permeability via both paracellular and transcellular pathways (Schepel et al., 2010; Lu et al.,

2011; see Beyenbach and Piermarini, 2011). The MTs of R. prolixus are composed of only

principal cells. With the absence of the stellate cells, the expression of the kinin receptor in MTs

is also likely absent, thus, the kinins have no direct diuretic activity in R. prolixus. Nonetheless,

the lack of Rhopr-kinins’ diuretic function has not affected the overall diuretic capabilities of R.

prolixus because this insect has other diuretic hormones. For example, serotonin is a primary

diuretic hormone in the kissing bug (Maddrell et al., 1991; see Orchard, 2001) with other

neuropeptides such as DH31 (Te Brugge et al., 2005) and Rhopr-CRF/DH (Te Brugge et al.,

2011) also involved in stimulating the secretion of the tubules in R. prolixus. Meanwhile, the

localization of the insect kinins to the R. prolixus midgut and hindgut as well as their

myostimulatory abilities indicates their crucial involvement in digestion and excretion.

Leucokinin I and CRF/DH-like peptides have both been co-localized to the PLNSCs of the

MTGM as well as in neurohaemal sites on ABNs 1 and 2 of R. prolixus (Te Brugge et al., 2001)

and such co-localization is also seen in the nervous systems of the tobacco hornworm (Chen et

al., 1994b) and locust (Thompson et al., 1995). In fact, the levels of both these neuropeptides

decrease from the Rhodnius CNS and neurohaemal sites 2.5 hours post feeding and are only

restored 24 hours after feeding (Te Brugge and Orchard, 2002b). Elsewhere, single-labelled cells

were observed for each neuropeptide family. Their co-localization and single localization thus,

signifies the possibility that these neuropeptides may either be released individually or together.

The theory of co-release in turn suggests that these neuropeptides could act on certain tissues

61

together, possibly through synergism. Synergism between kinin and CRF/DH has been observed

for MTs secretion of L. migratoria (Thompson et al., 1995) and M. domestica (Iboni et al.,

1998); however, synergism on the myotropic abilities of these peptides has not been examined

before. To test the theory of synergism on hindgut contractions, low doses of Rhopr-kinin 2 and

Rhopr-CRF/DH were tested on the hindgut. Independently, both peptides increase hindgut

contractions. In comparison, application of low doses of both peptides together on the hindgut

results in a greater increase in both the amplitude and frequency of contractions, although the

results are not statistically significant. Visually, the contractions were noted to be more robust

when both the peptides were applied on the hindgut. The differences in hindgut contractions

observed when the peptides are applied independently and then together also exemplify the

system’s flexibility. It is difficult to infer what increases in amplitude mean since the impedence

machine only records changes in contraction frequency. Upon co-application of the peptides,

frequency of contractions also increases, but is not statistically significant in comparison to

Rhopr-CRF/DH alone (Figure 7A and C). Concurrently, in cases where a reduction in the

contraction frequency is noted, it was compensated by a rise in the robustness of contractions.

We suggest that these peptides together enhance the myotropic activities of the hindgut of R.

prolixus in comparison to their independent myostimulatory effects. However, their combinative

effects do not appear to be synergistic. Previous studies investigating the second messenger

pathways employed by these peptides proposed that the kinins increase intracellular Ca2+ (Cady

and Hagedorn, 1999b; Radford et al., 2002; 2004; Pietrantonio et al., 2005), whereas the CRF-

like peptides appear to act via cAMP (see Te Brugge et al., 2009). In R. prolixus, Rhopr-

CRF/DH increase cAMP levels of the hindgut but leucokinins do not (see Orchard, 2009).

Accordingly, we hypothesize that the myotropic activities of these 2 peptides are a result of

independent signal transduction pathways, which lead to the enhanced myotropic activity. The

interactions of these two second messenger pathways are most likely responsible for eliciting the

strong hindgut contractions. For example, cross talk between second messenger pathways (i.e.

cAMP and inositol triphosphate-induced Ca2+) has been reported in blowfly salivary glands

(Schmidt et al., 2008) and a similar mechanism could be involved in this case.

In comparison to the effects of Rhopr-kinin 2 on the hindgut, both synthetic kinin analogs caused

a statistically significant increase in contraction (Figure 8A and B). Although the analogs

increased the amplitude of basal contractions, these effects were irreversible because the increase

62

in basal tonus could not be reversed despite numerous physiological saline washes (Figure 8Ai

and Bi). 1729[Φ1]wp-1 (Ac-R[Aib]FF[Aib]WGa) had a larger effect on the amplitude of

contractions compared to 1728[Φ1]wp-3 ([Aib]FF[Aib]WGa). This difference is presumably

associated with the composition of the kinin analogs. These analogs are synthetically devised

based on the notion that endogenous peptidases quickly degrade peptides such as kinins and

hence, their effect is short-lived on their target tissues (see Isaac et al., 2009). In particular, the

endopeptidase, angiotensin-converting enzyme (ACE) from the housefly has been shown to

inactivate the insect kinins through the hydrolysis of the dipeptide amide fragment at the C-

terminal (Nachman et al., 1997; 2011). The vulnerability of the insect kinins towards such

peptidases restricts their use as agents for pest management and thus, effective and biostable

kinins are being synthesized and tested. The substitution of the third amino acid residue with a

α,α-disubstituted amino acid, α-aminoisobutyric acid (Aib) has been shown to protect the peptide

from inactivation through steric hindrance (Nachman et al., 1997). In fact, the addition of a

second Aib at the N-terminus of the core pentapeptide motif adds further biostability to the

analog (see Nachman et al., 2009). Aib-kinin analogs have been shown in past studies to have

greater myotropic and diuretic capabilities in L. maderae and A. domesticus than the naturally

occurring kinins (Nachman et al., 1997). Thus, the Aib substitutions within the tested analogs

add stability to the structure of the kinins while maintaining their function. In fact, the

irreversibility of the basal tonus indicates that the synthetic kinin analogs were more tightly

bound to the kinin receptor(s) on the hindgut. The larger increase in amplitude of the phasic

contractions by 1729[Φ1]wp-1 when compared to 1728[Φ1]wp-3 may be attributed to the

acetylated arginine at the N-terminus of 1729[Φ1]wp-1 which is not present in analog

1728[Φ1]wp-3. However, since we were unable to test both these analogs on the same hindgut

preparation, it is impossible to confirm this. Different kinin analogs as well as ACE inhibitors

have previously been reported to inhibit weight gain in lepidopteran species like H. zea

(Nachman et al., 2003) and H. virescens (Seinsche et al., 2000). In the latter species, ACE

inhibitors have also been reported to significantly increase mortality rates and affect

development adversely. Therefore, such experiments demonstrate the potential for these kinds of

agonists to be used as biotechnological tools for agriculturally and medically-relevant pest

control in the future.

63

In totality, this study sheds light upon the molecular and physiological actions of the endogenous

insect kinins in the medically-important kissing bug. Studying the expression of the Rhopr-kinin

transcript as well as analyzing the myostimulatory activities of Rhopr-kinin peptides and their

cooperative involvement with other neuropeptides, have added to the knowledge of the feeding-

related physiological events which are responsible for Chagas’ disease transmission from this

insect to its vertebrate hosts. Advancing this research will provide a more comprehensive

understanding of the roles of insect kinins in digestion and excretion. Ultimately, this could lead

to the development of pharmacological tools that will prevent disease transmission from this

hemipteran to humans.

64

Future Directions

The initiation and completion of transcription of messenger RNA requires a promoter region and

a polyadenylation signal. Therefore, cloning of the 5’ and 3’ UTRs is essential in order to

confirm that the Rhopr-kinin cDNA does in fact translate into the expected prepropeptide

(although the presence of the peptides in the CNS extracts indicates it does). Multiple primer

designs will be designed and tested through temperature-gradient PCR before selecting the most

suitable ones for 5’ and 3’ RACE. Additionally, in situ hybridization will be performed on the

midgut to confirm previous immunohistological studies that the endocrine cells of the midgut

produce and process the Rhopr-kinin prepropeptide. This will further support the involvement of

insect kinins in digestion.

The molecular cloning and expression mapping of the Rhopr-kinin receptor(s) will add a breadth

of knowledge of the roles of Rhopr-kinins. Thus far, kinin receptors have been cloned from

A.aegypti (Pietrantonio et al., 2005), Anopheles stephensi (Radford, et al; 2004), D.

melanogaster (Radford et al., 2002), H. zea (Scherkenbeck, et al; 2009), in the Southern cattle

tick, Boophilus microplus (Holmes et al., 2000) and the pond snail L. stagnalis (Cox et al., 1997).

In all these species, a single kinin gene producing a single transcript variant has been isolated,

although the kinin receptors may undergo various tissue-specific post-translational modifications

(Radford et al., 2004; Kersch and Pietrantonio, 2011). Furthermore, in A. aegypti, the 3 Aedes

kinins are thought to act through the single endogenous kinin through a “ligand-dependent

selectivity of receptor signalling” where binding of different kinin isoforms to the receptor

initiates a different physiological response (Schepel et al., 2010). Thus, after analyzing the

cloning and spatial expression profile of the Rhopr-kinin receptor(s), it would be worth

functionally characterizing the receptors, especially since R. prolixus has a long list of

endogenous kinins. Transfecting the Rhopr-kinin receptor(s) into mammalian cell lines like the

Chinese Hamster Ovarian (CHO) cells will not only provide details on ligand-receptor binding,

but it will also enhance our understanding of secondary messenger pathways employed by

Rhopr-kinins to initiate signal transduction, so as to achieve flexibility in varying physiological

outcomes. Along with this, experiments designed specifically to understand the signal

transduction pathways of both Rhopr-kinins and Rhopr-CRF/DH are also important because they

65

will demonstrate how synergism is established in the MTs of the locust and the moth, but will

elucidate the enhanced hindgut contractions of R. prolixus upon co-application of the peptides.

Developmental differences and sexual dimorphisms in the expression of the Rhopr-kinins and

their receptor(s) as well as prandial and post-prandial changes in receptor expression studied

through real-time PCR and immunohistochemistry can enhance our understanding of the roles of

Rhopr-kinins and their receptor(s) in developmental, gender-specific and feeding-associated

events. Finally, the exploitation of the Rhopr-kinin transcript and its associated receptor

transcript(s) via RNA interference can also be performed to elucidate in vivo behavioural and

physiological roles of this family of neuropeptides as shown in Aedes (Kersch and Pietrantonio,

2011).

66

References

Al-Anzi, B., Armand, E., Nagamei, P., Olszewski, M., Sapin, V., Waters, C., Zinn, K.,

Wyman, R.J. and Bezer, S. (2010). The leucokinin pathway and its neurons regulate meal

size in Drosophila. Current Biology. 20, 969-978.

Alstein, M. and Nässel, D.R. (2010). Chapter 8: Neuropeptides signalling in insects. Advances

in experimental medicine and biology. 2010, 155-165.

Audsley, N., Goldsworthy, G.J. and Coast, G.M. (1997). Circulating levels of Locusta diuretic

hormone: the effect of feeding. 96, 59-65.

Blackburn, M.B., Wagner, R.M., Shabanowitz, J., Kochansky, J.P., Hunt, D.F. and Raina,

A.K. (1995). The isolation and identification of three diuretic kinins from the abdominal

ventral nerve cord of adult Helicoverpa zea. Journal of Insect Physiology. 41, 723-730.

Baldwin, D.C., Schegg, K.M., Furuya, K., Lehmberg, E. and Schooley, D.A. (2001).

Isolation and identification of a diuretic hormone from Zootermopsis nevadensis. Peptides.

22, 147-152.

Berlind, A. and Maddrell, S.H.P. (1979). Changes in hormone activity of single neurosecretory

cell bodies during a physiological secretion cycle. Brain Research. 161, 459-467.

Beyenbach K.W. and Piermarini, P.M. (2011). Transcellular and paracellular pathways of

transepithelial fluid secretion in Malpighian (renal) tubules of the yellow fever mosquito

Aedes aegypti. Acta Physiologica. 202, 387-407.

Cady, C and Hagedorn, H.H. (1999a). The effect of putative diuretic factors on in vivo urine

production in the mosquito, Aedes aegypti. Journal of Insect Physiology. 45, 317-325.

Cady, C and Hagedorn, H.H. (1999b). Effects of putative diuretic factors on intracellular

second messenger levels in the Malpighian tubules in Aedes aegypti. 45, 327-337.

67

Cantera, R., Hansson, B.S., Hallberg, E., N D.R. Nässel. (1992). Postembryonic development

of leucokinin I-immunoreactive neurons innervating a neurohemal organ in the turnip moth

Agrotis segetum. Cell & Tissue Research. 269, 65–77.

Cantera, R. and Nässel, D.R. (1992). Segmental peptidergic innervation of abdominal targets in

larval and adult dipteran insects revealed with an antiserum against leucokinin I. Cell &

Tissue Research. 269, 459–471.

Chen, Y., Veenstra, J.A. Davis, N.T. and Hagedorn, H.H. (1994a). A comparative study of

leucokinin-immunoreactive neurons in insects. Cell & Tissue Research. 276, 69-83.

Chen, Y., Veenstra, J.A. Davis, N.T. and Hagedorn, H.H. (1994b). Leucokinin and diuretic

hormone immunoreactivity of neurons in the tobacco hornworm, Manduca sexta, and co-

localization of this immunoreactivity in lateral neurosecretory cells of abdominal ganglia. Cell

& Tissue Research. 278, 493-507.

Claeys, I., Poels, J., Simonet, G., Franssens, V., Van Loy, T., Van Hiel, M.B., Breugelmans,

B. and Vanden Broeck, J. (2005). Insect neuropeptide and peptide hormone receptors:

current knowledge and future directions. Vitamins and Hormones. 73, 217-282.

Clottens, F.L., Meola, S.M., Coast, G.M., Hayes, T.K., Wright, M.S., Nachman, R.J. and

Holman, G.M. (1993). Characterization of an antiserum against an achetakinin I-analog and

its use for the localization of culekinin depolarizing peptide II in the mosquito, Culex

salinarius. Regulatory Peptides. 49, 145–157.

Coast, G. (2007). The endocrine control of salt balance in insects. General and Comparative

Endocrinology. (152), 332-338.

Coast, G. (2009). Neuroendocrine control of ionic homeostasis in blood-sucking insects. The

Journal of Experimental Biology. 212, 318-386.

Coast, G.M. and Garside, C.S. (2005). Neuropeptide control of fluid balance in insects. Annals

New York Academy of Sciences. 1040, 1-8.

68

Coast, G.M., Holman, G.M. and Nachman, R.J. (1990). The diuretic activity of a series of

cephalmyotropic neuropeptides, the achetakinins, on isolated Malpighian tubules of the house

cricket, Acheta domesticus. Journal of Insect Physiology. 36, 481-488.

Coast, G.M. and Schooley, D.A. (2011). Toward a consensus nomenclature for insect

neuropeptides and peptide hormones. Peptides. 32, 620-631.

Coast, G.M., Orchard, I., Phillips, J.E., and Schooley, D.A. (2002). Insect diuretic and

antidiuretic hormones. Advances in Insect Physiology. 29, 279-409.

Cox, K.J.A., Tensen, C.P., Van der Schors, R.C., Li, K.W., van Heerikhuizen, H.,

Vreugdenhil, E., Geraerts, W.P.M. and Burke, J.F. (1997). Cloning, characterization, and

expression of a G-protein-coupled receptor from Lymnaea stagnalis and identification of a

leucokinin-like peptide, PSFHSWSamide, as its endogenous ligand. The Journal of

Neuroscience. 17, 1197–1205.

Christie, A.E. (2008). In silico analyses of peptide paracrines/hormones in Aphidoidea. General

and Comparative Endocrinology. 159, 67-79.

Christie, A.E., Nolan, D.H., Garcia, Z.A., McCoole, M.D., Harmon, S.M., Congdon-Jones,

B., Ohno, P., Hartline, N., Congdon, C.B., Baer, K.N. and Lenz, P.H. (2011).

Bioinformatic prediction of arthropod/nematode-like peptides in non-arthropod, non-

nematode members of the Ecdysozoa. General and Comparative Endocrinology. 170, 480-

486.

Cook, B.J., Holman, G.M., Wagner, R.M. and Nachman, R.J. (1989). Pharmacological

actions of a new class of neuropeptides, the leucokinins I-IV, on the visceral muscles of

Leucopaea maderae. Comparative Biochemistry and Physiology. 93C, 257-262.

Cook, B.J., Holman, G.M., Wagner, R.M. and Nachman, R.J. (1990). Comparative

pharmacological actions of leucokinins V-VIII on the visceral muscles of Leucophaea

maderae. Comparative Biochemistry and Physiology. 95C, 19-24.

Dias, JCP., Silveira, AC. And Schofield, CJ. (2002). The impact of Chagas’ disease control in

Latin America – A review. Memórias do Instituto Oswaldo Cruz. 97, 603-612.

69

De Souza, W. (2002). Basic cell biology of Trypanosoma cruzi. Curr. Pharma. Des. 8, 269-285.

Edwards. J.S. (1998). Sir Vincent Wigglesworth and the coming of age of insect development.

International Journal of Developmental Biology. 42, 471-473.

Eigenheer, R.A., Nicolson, S.W., Schegg, K.M., Hull, J.J. and Schooley, D.A. (2002).

Identification of a potent antidiuretic factor acting on beetle Malpighian tubules. Proceedings

of the National Academy of Sciences of the United States of America. 99, 84-89.

Eigenheer, R.A., Wiehart, U.M., Nicolson, S.W., Schegg, K.M., Hull, J.J. and Schooley,

D.A. (2003). Isolation, identification and localization of a second beetle antidiuretic peptide.

Peptides. 24, 27-34.

Friend, W.G. and Smith, J.J.B. (1977). Factors affecting feeding by bloodsucking insects.

Annual Review of Entomology. 22, 309-331.

Gard, A.L., Lenz, P.H., Shaw, J.R. and Christie, A.E. (2009). Identification of putative

peptide paracrineséhormones in the water flea Daphnia pulex (Crustacea; Branchiopoda;

Cladocera) using transcriptomics and immunohistochemistry. General and Comparative

Endocrinology. 160, 271-287.

Gäde, G. (2004). Regulation of intermediary metabolism and water balance of insects by

neuropeptides. Annual Review of Entomology. 49, 93-113.

Gäde, G. and Goldsworthy G.J. (2003). Insect peptide hormones: a selective review of their

physiology and potential application for pest control. Pest Management Science. 59, 1063-

1075.

Gonzalez, R. and Orchard I. (2008). Characterization of neuropeptide F-like immunoreactivity

in the blood-feeding hemipteran, Rhodnius prolixus. Peptides. 29, 545-558.

Guillemin, R. (2011). Neuroendocrinology: a short historical review. Annals New York

Academy of Sciences. 1220, 1-5.

70

Hartenstein, V. (2006). The neuroendocrine system of invertebrates: a developmental and

evolutionary perspective. Journal of Endocrinology. 190, 555-570.

Hauser, F., Neupert, S., Williamson, M., Predel, R., Tanaka, Y., and Grimmelikhuijzen,

C.J.P. (2010). Genomics and peptidomics of neuropeptides and protein hormones in the

parasitic wasp Nasonia vitripennis. Journal of proteome research. 9, 5296-5310.

Hayes, T.K., Holman, G.M., Pannabecker, T.L., Wright, M.S., Strey, A.A., Nachman, R.J.,

Hoel, D.F., Olson, J.K. and Beyenbach, K.W. (1994). Culekinin depolarizing peptide: a

mosquito leucokinin-like peptide that influences insect Malpighian tubule ion transport.

Regulatory Peptides. 52, 235-248.

Holman, G.M., Cook, B.J. and Nachman, R.J. (1986a). Isolation, primary structure and

synthesis of two neuropeptides from Leucophaea maderae: members of a new family of

cephalomyotropins. Comparative Biochemistry & Physiology. 84C, 205–211.

Holman, G.M., Cook, B.J. and Nachman, R.J. (1986b). Primary structure and synthesis of two

additional neuropeptides from Leucophaea maderae: members of a new family of

cephalomyotropins. Comparative Biochemistry & Physiology. 84C, 271–276.

Holman, G.M., Cook, B.J. and Nachman, R.J. (1987a). Isolation, primary structure, and

synthesis of leucokinins V and VI: myotropic peptides of Leucophaea maderae. Comparative

Biochemistry & Physiology. 88C, 27–30.

Holman, G.M., Cook, B.J. and Nachman, R.J. (1987b). Isolation, primary structure and

synthesis of leucokinins VII and VIII: the final members of this new family of

cephalomyotropic peptides isolated from head extracts of Leucophaea maderae. Comparative

Biochemistry &Physiology. 88C, 31–34.

Holman, G.M., Nachman, R.J. and Coast, G.M. (1999). Isolation, characterization and

biological activity of a diuretic myokinin neuropeptide from the housefly, Musca domestica.

Peptides. 20, 1-10.

71

Holmes, S.P., He, H., Chen, A.C., Ivie, G.W. and Pietrantonio, P.V. (2000). Cloning and

transcriptional expression of a leucokinin-like peptide receptor from the Southern cattle tick,

Boophilus microplus (Acari: Ixodidae). Insect Molecular Biology. 9, 457-465.

Ianowski, J.P., Paluzzi, J.P., Te Brugge, V.A. and Orchard, I. (2010). The antidiuretic

neurohormone RhoprCAPA-2 downregulates fluid transport across the anterior midgut in the

blood-feeding insect Rhodnius prolixus. American Journal of Physiology. Regulatory,

Integrative and Comparative Physiology. 298, 548-557.

Iboni, A., Holman, G.M., Nachman, R.J., Orchard, I. and Coast, G.M. (1998).

Immunocytochemical localisation and biological activity of diuretic peptides in the housefly,

Musca domestica. Cell & Tissue Research. 294, 549-560.

Isaac, R.E., Bland, N.D. and Shirras, A.D. (2009). Neuropeptidases and the metabolic

inactivation of insect neuropeptides. General and Comparative Endocrinology. 162, 8-17.

Kataoka, H., Troetschler, R.G., Li, J.P., Kramer, S.J., Carney, R.L. and Schooley, D.A. (1989).

Isolation and identification of a diuretic hormone from the tobacoo hornworm, Manduca

sexta. Proceedings of the National Academy of Sciences of the United States of America. 86,

2976-2980.

Kay, I., Patel, M., Coast, G.M., Totty, N.F., Mallet, A.I. and Goldsworthy G.J. (1992).

Isolation, characterization and biological activity of a CRF-related diuretic peptide from

Periplanata Americana L. Regulatory Peptides. 42, 111-122.

Kersch, C.N. and Pietrantonio, P.V. (2011). Mosquito Aedes aegypti (L.) leucokinin receptor

is critical for in vivo fluid excretion post blood feeding. FEBS Letters. 585, 3507-3512.

Kim, Y-J., Žitňan, D., Cho, K-H., Schooley, D.A., Mizoguchi, A. and Adams, M.E. (2006).

Central peptidergic ensembles associated with organization of an innate behaviour.

Proceedings of the National Academy of Sciences of the United States of America. 103,

14211-14216.

Lane, N.J., Leslie, R.A. and Swales, L.S. (1975). Insect peripheral nerves: accessibility of

neurohaemal regions to lanthanum. Journal of Cell Science. 18, 179-197.

72

Lange, A.B. Orchard, I. and Barrett, F.M. (1989). Changes in haemolymph serotonin levels

associated with feeding in the blood-sucking bug, Rhodnius prolixus. Journal of Insect

Physiology. 35, 393-399.

Lee, B.H., Kang, H., Kwon, D., Park, C.I., Kim, W.K. and Kim, M.Y. (1998). Postembryonic

development of leucokinin-like immunoreactive neurons in the moth Spodopter litura. Tissue

& Cell. 30, 74-85.

Li, B., Predel, R., Neupert, S., Hauser, F., Tanaka, Y., Cazzamali, G., Williamson, M.,

Arakane, Y., Verleyen, P., Schoofs, L., Schachtner, J., Grimmelikhuijzen, C.J.P. and

Park, Y. (2008). Genomics, transcriptomics and peptidomics of neuropeptides and protein

hormones in the red flour beetle Tribolium castaneum. Genome Research. 18, 113-122.

Lima, F.M., Oliveira, P., Mortara, R.A., Silveira, J.F. and Bahia, D. (2010). The challenge of

Chagas’ disease: Has the human pathogen, Trypanosoma cruzi, learned how to modulate

signalling events to subvert host cells? New Biotechology. 27, 837-843.

Lovejoy, D.A. and Barsyte-Lovejoy, D. (2010). Characterization of a corticotrophin-releasing

factor (CRF)/diuretic hormone-like peptide from tunicates: Insight into the origins of the

vertebrate CRF family. General and Comparative Endocrinology. 165, 330-336.

Lovejoy, D.A. and Jahan, S. (2006). Phylogeny of the corticotrophin-releasing factor family of

peptides in the metazoa. General and Comparative Endocrinology. 146, 1-8.

López-Arias, B., Dorado, B., and Herrero, P. (2011). Blockade of the release of the

neuropeptide leucokinin to determine its possible functions in fly behaviour: chemoreception

assays. Peptides. 32, 545-552.

Lu, H.L., Kersch, C. and Pietrantonio, P.V. (2010). The kinin receptor is expressed in the

Malpighian tubule stellate cells in the mosquito Aedes aegypti (L.): A new model needed to

explain ion transport? Insect Biochemistry and Molecular Biology. 41, 135-140.

73

Maddrell, S.H.P. (1963). Excretion in the blood-sucking bug, Rhodnius prolixus Stål. I. The

control of diuresis. The Journal of Experimental Biology.40, 247-256.

Maddrell, S.H.P. (1964a). Excretion in the blood-sucking bug, Rhodnius prolixus Stål. II. The

control of diuresis and the effect of temperature. The Journal of Experimental Biology.41,

163-176.

Maddrell, S.H.P. (1964b). Excretion in the blood-sucking bug, Rhodnius prolixus Stål. III. The

control of the release of the diuretic hormone. The Journal of Experimental Biology.41, 459-

472.

Maddrell, S.H.P. (1966a). The site of release of the diuretic hormone in Rhodnius—A new

neurohaemal system in insects. The Journal of Experimental Biology. 45, 499-508.

Maddrell, S.H.P. (1966b). Nervous control of the mechanical properties of the abdominal wall

at feeding in Rhodnius. The Journal of Experimental Biology. 44, 59-68.

Maddrell, S.H.P. (1969). Secretion by the Malpighian tubules of Rhodnius. The movements of

ions and water. The Journal of Experimental Biology. 51, 71-97.

Maddrell, S.H.P., Herman, W.S., Mooney, R.L. and Overton, J.A. (1991). 5-

Hydroxytryptamine: A second diuretic hormone in Rhodnius prolixus. The Journal of

Experimental Biology. 156, 557-566.

Maddrell, S.H.P. and Phillips, J.E. (1975). Secretion of hypo-osmotic fluid by the lower

Malpighian tubues of Rhodnius prolixus. The Journal of Experimental Biology. 62, 671-683.

Massaro, R.C., Lee, L.W., Patel, A.B., Wu, D.S., Yu, M.J., Scott, B.N., Schooley, D.A.,

Schegg, K.M. and Beyenbach, K.W. (2004). The mechanism of action of the antidiuretic

peptide Tenmo ADFa in Malpighian tubules of Aedes aegypti. The Journal of Experimental

Biology. 207, 2877-2888.

Meeusen, T., Mertens, I., De Loof, A. and Schoofs, L. (2003). G Protein-Coupled Receptors in

invertebrates: a state of the art. International Review of Cytology. 230, 189-261.

74

Nachman, R.J., Coast, G.M., Holman, G.M. and Beier, R.C. (1995). Diuretic activity of C-

terminal group analogues of the insect kinins in Acheta domesticus. Peptides. 16, 809-813.

Nachman, R.J., Coast, G.M., Douat, C., Fehrentz, J-A., Kaczmarck, K., Zabrocki, J.,

Pryor, N.W. and Martinez, J. (2003). A C-terminal aldehyde insect kinin analog enhances

inhibition of weight gain and induces significant mortality in Helicoverpa zea larvae.

Peptides. 24, 1615-1621.

Nachman, R.J., Isaac, R.E., Coast, G.M. and Holman, G.M. (1997). Aib-containing

analogues of the insect kinin neuropeptide family demonstrate resistance to an insect

angiotensin-converting enzyme and potent diuretic activity. Peptides. 18, 53-57.

Nachman, R.J., Mahdian, K, Nässel, D.R., Isaac, R.E., Pryor, N. and Smagghe, G. (2011).

Biostable multi-Aib analogs of tachykinin-related peptides demonstrate potent oral aphicidal

activity in the pea aphid Acyrthosiphon pisum (Hemiptera: Aphidae). Peptides. 32, 587-594.

Nachman, R.J., Pietrantonio, P.V. and Coast, G.M. (2009). Toward the development of novel

pest management agents based upon insect kinin neuropeptide analogues. Annals of the New

York Academy of Sciences. 1163, 251-261.

Nässel, D.R. (2002). Neuropeptides in the nervous system of Drosophila and other insects:

multiple roles as neuromodulators and neurohormones. Progress in Neurobiology. 68, 1-84.

Nässel, D.R. (2009). Neuropeptide signalling near and far: how localized and timed is the action

of neuropeptides in brain circuits? Invertebrate Neuroscience. 9, 57-75.

Nässel, D.R., Cantera, R. and Karlsson, A. (1992). Neurons in the cockroach nervous system

reacting with antisera to the neuropeptide leucokinin I. The Journal of Comparative

Neurology. 322, 45-67.

Nieto, J., Veelaert, D., Derua, R., Waelkens, E., Cerstiaens, A., Coast, G., Devreese, B., Van

Beeumen, J., Calderon, J., De Loof, A., and Schoofs, L. (1998). Identificationof one

tachykinin- and two kinin-related peptides in the brain of the white shrimp, Penaeus

vannamei. Biochemical and Biophysical Research Communications. 248, 406–411.

75

Nieto, J., Veenstra, J., Waelkens, E., Calderon, J., Baggerman, G., Veelaert D., De Loof, A.,

and Schoofs, L. (1999). Isolation and identification of one tachykinin and three kinin-related

peptides in the central nervous system of Penaeus vannamei. In Proceedings of the Fourth

International crustacean Congress. F.R. Schram & J.C. von Vaupel Klein, Eds. p. 941-950.

Brill. Leiden, the Netherlands.

O’Donnell, M.J., Reault, M.R., Davies, S.A., Rosay, P., Harvey, B.J., Maddrell, S.H.P.,

Kaiser, K. and Dow, J.A. (1998). Hormonally controlled chloride movement across

Drosophila tubules is via ion channels in stellate cells. American Journal of Physiology.

Regulatory, Integrative and Comparative Physiology. 274, R1039-R1049.

O’Donnell, M.J. (2009). Too much of a good thing: how insects cope with excess ions or toxins

in the diet. The Journal of Experimental Biology. 212, 363-372.

Oeh, U. and Nauen, A.R. (2003). Myotropic effect of helicokinins, tachykinin-related peptides

and Manduca sexta allatotropin on the gut of Heliothis virescens (Lepidoptera: Noctuidae).

Journal of Insect Physiology. 49, 323-337.

Ons, S., Richter, F., Urlaub, H. and Rivera-Pomar, R. (2009). The neuropeptidome of

Rhodnius prolixus brain. Proteomics. 9, 788-792.

Ons, S., Sterkel, M., Diambra, L., Urlaub, H. and Rivera-Pomar, R. (2011). Neuropeptide

precursor gene discovery in the Chagas disease vector Rhodnius prolixus. Insect Molecular

Biology. 20, 29-44.

Orchard, I. (2006). Serotonin: A coordinator of feeding-related physiological events in the

blood-gorging bug, Rhodnius prolixus. Comparative Biochemistry & Physiology, Part A. 144,

316-324.

Orchard, I. (2009). Peptides and serotonin control feeding-related events in Rhodnius prolixus.

Frontiers in Bioscience. E1, 250-262.

Orchard, I. and Te Brugge, V. (2002). Contractions associated with the salivary glands of the

blood-feeding bug, Rhodnius prolixus: evidence for both a neural and neurohormonal

coordination. Peptides. 23, 693-700.

76

Orchard, I., Lange, A.B., and Bendena, W.G. (2001). FMRFamide-related peptides:

Multifunctional family of structurally related neuropeptides in insects. Advances in insect

physiology. 28, 267-329.

Paluzzi, J.P., Russell, W.K., Nachman, J. and Orchard, I. (2008). Isolation, Cloning and

Expression Mapping of a Gene Encoding an Antidiuretic Hormone and Other CAPA-Related

Peptides in the Disease Vector, Rhodnius prolixus. Endocrinology. 149, 4638-4646.

Patel, M., Chung, J-S., Kay, I., Mallet, A.I., Gibbon, C.R., Thompson, K.S.J., Bacon, J.P.

and Coast, G.M. (1994). Localization of Locusta-DP in locust CNS and haemolymph

satisfies initial hormonal criteria. Peptides. 15, 591-602.

Patel, M., Hayes, T.K. and Coast, G.M. (1995). Evidence for the hormonal function of a CRF-

related diuretic peptide (Locusta-DP) in Locusta migratoria. The Journal of Experimental

Biology. 198, 793-804.

Petersen, T.N., Brunak, S., von Heijne, G. and Nielsen, H. (2011). SignalP4.0: discriminating

signal peptides from transmembrane regions. Nature methods. . 8, 785-786.

Pietrantonio, P.V., Jagge, C., Taheja-Bageshwar, S., Nachman, R.J. and Barhoumi, R.

(2005). The mosquito Aedes aegypti (L.) leucokinin receptor is a multiligand receptor for the

three Aedes kinins. Insect Molecular Biology. 14, 55-67.

Predel, R., Kellner, R., Rapus, J., Penzlin, H. and Gäde, G. (1997). Isolation and structural

elucidation of eight kinins from the retrocerebral complex of the American cockroach,

Periplaneta americana. Regulatory Peptides. 71, 199-205.

Radford, J.C., Davies, S.A. and Dow, J.A. (2002). Systematic G-protein-coupled receptor

analysis in Drosophila melanogaster identifies a leucokinin receptor with novel roles. The

Journal of Biological Chemistry. 277, 38810-38817.

Radford, J.H., Terhzaz, S., Cabrero, P., Davies, S-A. and Dow, J.A.T. (2004). Functional

characterization of the Anopheles leucokinins and their cognate G-protein coupled receptor.

The Journal of Experimental Biology. 207, 4573-4586.

77

Reese M.G. (2001). Application of a time-delay neural network to promoter annotation in the

Drosophila melanogaster genome. Computers & Chemistry. 26, 51-56.

Reese M.G., Eeckman, F.H., Kulp, D. and Haussler, D. (1997). Improved splice site detection

in genie. Journal of Comparative Biology. 4, 311-323.

Roberts, V.A., Nachman, R.J., Coast, G.M., Hariharan, M., Chung, J.S., Holman, G.M.,

Williams, H. and Tainer, J.A. (1997). Consensus chemistry and β-turn conformation of the

active core of the insect kinin neuropeptide family. Chemistry & Biology. 4, 105-117.

Roller, L., Yamanaka, N., Watanabe, K., Daubnerová. Žitňan, D., Kataoka, H. and

Tanaka, Y. (2008). The unique evolution of neuropeptide genes in the silkworm Bombyx

mori. 38, 1147-1157.

Scharrer, B. (1987). Insects as models in neuroendocrine research. Annual Review of

Entomology. 32, 1-16.

Schepel, S.A., Fox, A.J., Miyauchi, J.T., Sou, T., Yang, J.D., Lau, K., Blum, A.W.,

Nicholson, L.K., Tiburcy, F., Nachman, R.J., Piermarini, P.M. and Beyenbach, K.W.

(2010). The single kinin receptor signals to separate and independent physiological pathways

in Malpighian tubules of the yellow fever mosquito. American Journal of Physiology.

Regulatory, Integrative and Comparative Physiology. 299, 612-622.

Scherkenbeck, J., Antonicek, H.-P., Vogelsang, K., Zdobinsky, T., Brücher, K., Rehländer,

D. and Chen, H. (2009). Receptor assay guided structure-activity studies of helicokinin insect

neuropeptides and peptidomimetic analogues. Journal of Peptide Science. 15, 783–789.

Schmid A. and Becherer, C. (1996). Leucokinin-like immunoreactive neurones in the central

nervous system of the spide Cupiennius salei. Cell & Tissue Research. 284, 143-152.

Schmidt, R., Baumann, O., and Walz, B. (2008). cAMP Potentiates InsP3-induced Ca2+

Release from the Endoplasmic Reticulum in Blowfly Salivary Glands. BMC Physiology. 8, 1-

11.

78

Schoofs, L, Holman, G.M., Proost, P., Van Damme, J., Hayes, T.K. and De Loof, A. (1992).

Locustakinin, a novel myotropic peptide from Locusta migratoria, isolation, primary structure

and synthesis. Regulatory Peptides. 37, 49-57.

Seinsche, A., Dyker, H., Lösel, P., Backhaus, D. and Scherkenbeck, J. (2000). Effects of

helicokinins and ACE inhibitors on water balance and development of Heliothis virescens

larvae. Journal of Insect Physiology. 46, 1423-1431.

Smart, D., Johnston, C.S., Halton, D.W. and Buchanan, K.D. (1993). Use of specific antisera

for the localisation and quantification of leucokinin immunoreactivity in the nematode,

Ascaris suum. Comparative Biochemistry & Physiology. 106C, 517-522.

Subramanian, P. (1994). Localization of leucokinin-like immunoreactive neurons and their

metamorphic changes in the ventral ganglion of the fly, Sacrophaga bullata. Comparative

Biochemistry & Physiology. 109A, 151-155.

Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S. (2011). MEGA5:

molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance,

and maximum parsimony methods. Molecular Biology and Evolution. 28, 2731-2739.

Te Brugge, V.A., Ianowski, J.P. and Orchard, I. (2009). Biological activity of diuretic factors

on the anterior midgut of the blood-feeding bug, Rhodnius prolixus. General and

Comparative Endocrinology. 162, 105-112.

Te Brugge, V.A., Lombardi, V.C., Schooley, D.A. and Orchard, I. (2005). Presence and

activity of a Dippu-DH31-like peptide in the blood-feeding bug, Rhodnius prolixus. Peptides.

26, 29-42.

Te Brugge, V.A., Miksys, S.M., Coast, G.M., Schooley, D.A. and Orchard, I. (1999). The

distribution of a CRF-like diuretic peptide in the blood-feeding bug Rhodnius prolixus. The

Journal of Experimental Biology. 202, 2017-2027.

Te Brugge, V.A., Nässel, D.R., Coast, G.M., Schooley, D.A. and Orchard, I. (2001). The

distribution of a kinin-like peptide and its co-localization with a CRF-like peptide in the

blood-feeding bug, Rhodnius prolixus. Peptides. 22, 161-173.

79

Te Brugge, V.A., Paluzzi, J.P., Neupert, S., Nachman, R.J., Orchard, I. (2010). Identification

of kinin-related peptides in the disease vector, Rhodnius prolixus. Peptides. 32, 469-474.

Te Brugge, V.A., Paluzzi, Schooley, D.A. and Orchard, I. (2011). Identification of the elusive

peptidergic diuretic hormone in the blood-feeding bug Rhodnius prolixus: a CRF-related

peptide. The Journal of Experimental Biology. 214, 371-381.

Te Brugge, V.A., Schooley, D.A. and Orchard, I. (2002a). The Biological Activity of Diuretic

Factors in Rhodnius prolixus. Peptides. 23, 671-681.

Te Brugge, V.A. and Orchard, I. (2002b). Evidence of CRF-like and kinin-like peptides as

neurohormones in the blood-feeding bug, Rhodnius prolixus. Peptides. 23, 1967-1979.

Terhzaz, S., O’Connell, F.C., Pollock, V.P., Kean, L., Davies, S.A., Veenstra, J.A. and Dow,

J.A.T. (1999). Isolation and characterization of a leucokinin-like peptide of Drosophila

melanogaster. The Journal of Experimental Biology. 202, 3667-3676.

Thompson, K.S.J., Rayne, R.C., Gibbon, C.R., May, S.T., Patel, M., Coast, G.M. and

Bacon, J.P. (1995). Cellular colocalization of diuretic peptides in locusts: a potent control

mechanism. Peptides. 16, 95–104.

Torfs, P., Nieto, J., Veelaert, D., Boon, D., Van De Water, G., Waelkens, E., Derua, R.,

Calderon, J., De Loof, A., and Schoofs, L. (1999). The kinin peptide family in invertebrates.

Annals of the Ney York Academy of Sciences. 897, 361-373.

Tsang, P.W. and Orchard, I. (1991). Distribution of FMRFamide-related peptides in the blood-

feeding bug, Rhodnius prolixus. The Journal of Comparative Neurology. 311, 17-32.

Veenstra, J.A. (1994). Isolation and identification of three leucokinins from the mosquito Aedes

aegypti. Biochemical and Biophysical Research Communications. 202, 715-719.

Veenstra, J.A. (2000). Mono- and Dibasic proteolytic cleavage sites in insect neuroendocrine

peptide precursors. Archives of Insect Biochemistry and Physiology. 43, 49-63.

80

Veenstra, J.A., Pattillo, J.M. and Petzel, D.H. (1997). A single cDNA encodes all three Aedes

leucokinins, which stimulate both fluid secretion by the Malpighian tubules and hindgut

contractions. The Journal of Biological Chemistry. 272, 10402-10407.

Veenstra, J.A., Rodriguez, L. and Weaver, R.J. (2012). Allatotropin, leucokinin and AKH in

honey bees and other hymenoptera. Peptides. DOI:

http://dx.doi.org.myaccess.library.utoronto.ca/10.1016/j.peptides.2012.02.019.

World Health Organization (WHO). (2008). Chagas disease: control and elimination—Report

of the Secretariat. Executive Board, Session 124th. Provisional agenda item 4.14, 1-4.

Žitňan, D., Kingan, T.G., Hermesman, J.L. and Adams, M.E. (1996). Identification of

ecydsis-triggering hormone from an epitracheal endocrine system. Science. 271, 88-91.

81

Appendix:

Molecular Identification and Characterization of Two Putative Serotonin Receptors in the

Kissing Bug, Rhodnius prolixus

82

Abstract

Rhodnius prolixus is a blood-feeding insect that transmits the parasitic protozoan, Trypanosoma

cruzi, which causes Chagas’ disease in humans. After a blood meal, this insect undergoes a rapid

diuresis eliminating excess water and salts from the blood meal that is regulated by diuretic

hormones. Serotonin or 5-hydroxytryptamine (5-HT) is a biogenic amine that plays a crucial role

as a neurotransmitter and a neurohormone, to modulate diuretic, digestive and feeding activities

of this insect. For example, 5-HT is involved in stimulating fluid secretion by Malpighian tubules

(MTs) following a dramatic rise in haemolymph 5-HT levels upon feeding. Based on previous

physiological and pharmacological studies, a vertebrate 5-HT type-2 (5-HT2)-like receptor is

believed to play an essential role in the post-prandial diuresis. In this study, we identify and

characterize a 5-HT2 receptor-like gene using a MT cDNA library. A partial sequence encoding a

5-HT2 receptor-like protein was identified; however, quantification of this putative receptor

revealed expression was inconsistent with a role in diuresis. Next, using a Drosophila mojavensis

5-HT2 receptor-like sequence as query, new primers were designed based on hits in the R.

prolixus preliminary genome assembly. A sequence has been amplified with high transcript

levels in the MTs, central nervous system, foregut and salivary glands. This gene is predicted to

be a classical G protein-coupled receptor with 7 transmembrane domains. So far, an incomplete

nucleotide sequence encoding 7 transmembrane domains has been isolated and the 5’end along

with the 5’ and 3’ untranslated regions of the transcript are still missing. Ongoing research will

confirm if this is the 5-HT receptor of interest.

83

Introduction

The medically-relevant blood-feeding hemipteran, Rhodnius prolixus, is one of 12 Triatominae

species known to transmit the parasitic protozoan Trypanosoma cruzi, and so causing Chagas’

disease in humans. Also known as the kissing bug, R. prolixus prevalently inhabits central and

South America where it infects approximately 16-18 million people with T. cruzi and results in

50, 000 deaths annually (Lima et al., 2010). Fifth instar R. prolixus in particular, consume large

blood meals that are 10 times their body weight and subsequently loose 40% of the weight of

their blood meal through rapid diuresis, thus, also making them interesting organisms to study

ion and osmoregulation (see Orchard et al., 2006). The obvious threat imposed by these dramatic

feeding habits is the restriction of mobility which in turn subjects the insect to being detected by

the host or predation. However, on a physiological and evolutionary standpoint, the consumption

of a large volume of mammalian, hypo-osmotic blood can endanger the haemolymph osmolarity

of the insect. Therefore, to maintain homeostasis, this insect removes excess ions and water in

the form of urine, sometimes even while it is feeding (see Orchard 2005; 2009). If the parasite-

containing urine is excreted onto the host, it may enter the victim through any open wound or

mucus membranes (e.g. eyes) and then travel into their bloodstream (WHO). Thus, rapid diuresis

and its regulating factors constitute an imperative area of research in the field of insect

physiology.

The rapid, post-prandial diuresis in post-embryonic stages of R. prolixus is regulated via various

neurohormonal pathways, comprising of both diuretic and anti-diuretic agents (see Coast, 2007

and Orchard, 2009). Diuretic factors like the biogenic (indole) amine, serotonin or 5-

hydroxytryptamine (5-HT) and peptides like the corticotropin-releasing factor (CRF)-like, kinin-

like, and calcitonin (CT)-like peptides are involved in increasing secretion of primary urine by

the Malpighian tubules (MTs) whereas the cardioactive peptide 2b (CAP2b), is known to induce

anti-diuretic effects on this insect’s system (see Coast 2007; Paluzzi et al., 2008). These diuretic

peptides may act on their target tissue either synergistically or additively (see O’Donnell and

Spring, 2000). For example, the Locusta-diuretic hormone (DH) and leucokinin have been shown

to act synergistically (Thompson et al., 1995) on the secretions of the locust MTs, and serotonin

and Rhopr-CRF/DH are also synergistic in R. prolixus (Paluzzi et al., 2012).

84

Of particular interest, is serotonin, because it serves as a neurotransmitter and/or a neurohormone

and plays a crucial role in modulating diuresis, digestion and other feeding activities of this

insect (see Orchard, 2009). Interestingly, serotonin is a key modulator of feeding-related

behaviours (e.g. salivation, swimming, contractions of the pharynx, etc ;) in another blood-

feeding invertebrate, the medicinal leech (Lent and Dickinson, 1988). In insects, serotonin-

related immunohistochemistry has been studied in several dipteran, orthopteran, lepidopteran,

blattarian species (see Nässel, 1988). As seen in vertebrates, serotonin also regulates diverse

physiological and behavioural processes in insects which include feeding-associated events like

secretory activities of the salivary glands in the locust (Ali et al., 1993); blowfly (Heslop and

Berridge, 1980) and a cockroach (Troppmann et al., 2007), muscle relaxation in the locust

(Molaei and Lange, 2003) as well as a decrease in feeding in an ant (Falibene et al., 2012).

Extensive studies in Drosophila melanogaster and Apis mellifera have linked 5-HT to

development and a string of behaviours associated with central pattern generators that control

olfactory learning, memory and circadian rhythms (Johnson et al., 2011; see Blenau and Thamm,

2011).

Serotonin was first discovered as a diuretic factor in R. prolixus by Simon Maddrell and

colleagues (1991). The first initial 5 minutes of feeding dramatically stimulates the elevation of

haemolymph levels of serotonin from 7nM to 115nM (Lange et al., 1989). Serotonin has been

shown to co-localize with the DH31 peptide in the dorsal unpaired medial (DUM) cells of the

MTGM and neurohaemal sites of all 5 abdominal nerves, therefore suggesting their potential for

co-release (Orchard et al., 1989 and Te Brugge et al., 2005). As the primary diuretic factor in R.

prolixus 5-HT stimulates muscle contractions of the dorsal vessel and hindgut, oesophagus

(foregut), salivary glands, crop, and is also associated with the plasticization of the cuticle and

the expulsion of waste (Ochard et al., 1988 and see Orchard, 2006). Serotonin-like

immunoreactivity is observed over these regions since they are innervated by serotonergic nerves

that supply 5-HT to these locations (Lange et al., 1989; see Orchard et al., 2006).

Additionally, as a diuretic factor, 5-HT is involved in the production of primary urine; by

stimulating secretion of excess water and ions by the Malpighian tubules (MTs) (see Martini et

al., 2007). The distal tubules reabsorb water and ions (e.g. Na+, K+, Cl-) whereas the proximal

tubule secrete K+ and Cl-. This is accomplished through numerous ion transporters and aquaporin

85

channels found in the tubules (see Martini et al., 2007 and O’Donnell, 2009). Unlike other organs

of R. prolixus, the MTs lack innervation and are thus affected by the drastic rise in the

haemolymph levels of serotonin (Lange et al., 1989). As a result, the activity of the tubules in

primary urine secretion is dependent on the presence of serotonin receptors in the MTs epithelial

cells (i.e. principal cells). Activation of the serotonin receptors results in increased activity of ion

and water transport channels that augment secretion in the distal tubules and increase

reabsorption of ions in the proximal tubule through a cAMP-mediated signal transduction

pathway (see Coast, 2007 and Martini et al., 2007).

Research studies have thus far predicted three known classes of serotonin receptors in several

insects: 5-HT receptors type 1, type 2 and type 7 (see Tierney, 2001). In comparison, 7 different

classes have been examined in vertebrates with varying modes of signal transduction (see

Raymond et al., 2001). Serotonin receptors have been predicted and/or cloned from several

insects and their endogeneous roles are now being investigated (e.g. Troppman et al., 2010;

Thamm et al., 2010; Dacks et al., 2006; von Nickisch-Rosenegk et al., 1996). However, the

knowledge of serotonin receptors in hemipterans like R. prolixus is little to none and has to be

explored in further depths.

Thus, the purpose of this research study was to clone and sequence the cDNA encoding the

serotonin receptor from the MTs of 5th instar R. prolixus. Upon isolation and identification of the

serotonin receptor gene, the objective of the study was to characterize and localize the serotonin

receptor and to study its spatial expression profile. Studying the endogenous serotonin receptors

in this disease vector will not only enhance the understanding of serotonin’s role in regulating the

post-prandial diuresis and feeding activities of R. prolixus but may also benefit pharmaceutical

industries interested in preventing Chagas’ disease through the inhibition or reduction of diuretic

rates of this hemipteran.

86

Materials & Methods

Animals Used

In this study, 5th instar Rhodnius prolixus Stål were raised at 25°C in incubators at a relatively

high humidity. The insect lines were fed regularly on rabbits’ blood. When dissecting to retrieve

different tissues, the insects were bathed in nuclease-free physiological saline phosphate-buffered

saline (PBS) (Sigma-Aldrich, Oakville, ON, Canada). When fed R. prolixus were needed for

dissections, they were fed on an artificial diet of saline with ATP (which triggers the release of 5-

HT) and not rabbits’ blood in order to avoid contamination with mammalian RNA. Dissected

tissues were then stored in RNAlater™ RNA stabilization reagent (Qiagen Inc., Mississauga,

ON, Canada).

Analysis of a Partially Cloned Putative Rhopr5-HT2 Receptor Gene

In the summer of 2009, an undergraduate research student, Song Kim worked on the molecular

cloning of a putative 5-HT type 2 receptor (5-HT2) gene. She was able to clone and sequence

portions of the receptor gene through a process of screening a MTs cDNA library. Starting from

where she left the project, all her cloning and sequencing results were first confirmed using pre-

existing primers (Table 1). All cloning procedures followed are described in the section below.

Next, a Contig, named SK_Rhopr5-HT2R, was built and the ORF encoded by the consensus

sequence was analyzed via protein BLAST on ExPASy (http://ca.expasy.org/tools/dna.html) to

compare it with other known insect 5-HT receptors. Since the receptor gene Contig was

amplified in segments previously, primers were generated to amplify the entire cDNA as well as

to amplify the 5’ and 3’ UTRs. Finally, Reverse Transcription-Polymerase Chain Reaction (RT-

PCR) was performed to understand spatial expression of the receptor transcript as described in

below sections. Any post-prandial changes in the expression of the receptor transcript were also

studied through RT-PCR.

87

Table 1: A list of the primer designs that were used to amplify partial segments of the predicted 5-HT2 receptor. The partial sequences were used to generate the Contig: SK_Rhopr5-HT2R. Oligo-nucleotide name Oligo-nucleotide sequence (5’-3’) Jpp5HTR2-F1 TCGTTATATATGGAGTGGGAGAATACC Jpp5HTR2-F2 TTCTGTTCGTATTGTTGTTCGTAGC Jpp5HTR2-R1 ATGACTGGAGTGAAGATCTTAATGG Jpp5HTR2-R2 TGTCCCTAGGTTTGTTGTTCG 5HTR2 349-424F1 GGTGCAATATATACGTAACATGTGATG 5HTR2 349-424F2 TGCATTTCATTAGGAAGATATCTTGG 5HTR2 349-424R1 TCCAAGATATCTTCCTAATGAAATGC 5HTR2 349-424R2 CATCACATGTTACGTATATATTGCACC 5HTR2 430-495F1 GGATCTTTGGTTCATTGGTAGC 5HTR2 430-495F2 ATGGTAATCATGGTAGTGACGTACG 5HTR2 430-495R1 AAACGGCCGCCTAATCTTC 5HTR2 430-495R2 AAACTGTCGCTCTTCTTCTTCG

A Second Attempt: In Silico Searches and Preliminary Molecular Cloning of a Putative

Rhopr5-HT2 Receptor Gene

Schaerlinger and colleagues have previously studied the 5-HT2 receptor in D. melanogaster

(2007). The amino acid and nucleotide sequences of the serotonin receptor type 2, isoform B was

retrieved from the NCBI website (GenBank Ac: NM_169011). These sequences were uploaded

onto the Geneious software 4.6.1 (Biomatters, Ltd., Aukland, New Zealand). The protein

sequence of D. melanogaster 5-HT2B receptor was used as a query sequence for tBLASTn

against the preliminary Rhodnius genome database. Primers were then created over regions that

had the highest similarities with the query sequence (Table 2).

Once these primers (5htr2-51.1for1 and 5htr2-51.1rev1) were tested via PCR, the preliminary

sequencing results were used as BLAST query sequence on the ExPASy web portal. The BLAST

hits revealed that the sequencing results were most similar to an un-annotated sequence from

Drosophila mojavensis but shared similarities with 5-HT2 receptors in other species (Ac:

B4KCQ3). Thus, the D. mojavensis protein sequence was used as query for tBLASTn against the

preliminary Rhodnius genome database. Once again, the highest BLAST hits were obtained and

gene-specific primers were generated on potential exon regions in order to amplify a partial

cDNA sequence (Table 2). All primers were created using Sigma Aldrich OligoCalculator to

design optimal primers.

88

Table 2: A table displaying the primer designs that were created over the tBLASTn hits between the D. melanogaster, D. mojavensis and the preliminary genome database of R. prolixus in order

to clone GB_Rhopr5-HT2R. Exon Oligo-nucleotide name Oligo-nucleotide sequence (5’-3’)

6

5htr2-51.1for1 TCGCGCACTTCATCTCG 5htr2-51.1rev1 TTCTTGAATGCTTGTCGAAAC SR2 58.0F1 ACGTGTCCCTATTTTGGTG SR2 58.0F2 CAAATACTATAGTAACATGGGATTCC SR2 58.0R1 TGTTATTGTTACATCTGCCTACG SR2 58.0R2 CGATATCGACACAATAAGACTTTC

5 SR2 60.0F1 GTTTGGAAAGACGAGGTACTTG SR2 60.0R1 TGCACCAAACTGTTGTAAAGC

4 SR2 73.0F1 CAGGATTACAACTCATTGGTGG

SR2 73.0R1 CGCTCGCCCAACC

3 SR2 87.3F1 AGTTTACTGTTTGGCGTGG SR2 87.3R1 CTAAGTGGAAGACTCATAGCTATCG

2 5htr2-58.2for1 GCAGCTGGCAACATCC 5htr2-58.2rev1 GCATGACGAGTATGGCGAC

Molecular Cloning of the Rhopr5-HT2 Receptor

A partial sequence obtained for this newly predicted 5-HT receptor (named GB_Rhopr5-HT2R)

using the 5htr2-51.1for1 and 5htr2-51.1rev1 primer set. In order to extend the upstream and

downstream regions of the partial serotonin receptor, a pre-made 5th instar MTs cDNA library

was screened (Table 2). This cDNA library was synthesized by Jean-Paul Paluzzi (Paluzzi et al.,

2008) using the Creator™ SMART™ cDNA Library Construction kit (ClonTech, Mountain

View, CA, USA). Each reaction comprised of the ThermoPol Buffer, Taq polymerase (New

England BioLabs, Pickering, ON, Canada), a vector-specific primer, a gene-specific primer,

dNTP mix and the template. The temperature for all reactions were programmed in the ®PCR

System 9700 thermal cycler (PE Applied Biosystems, Carlsbad, CA, USA). The temperature

program was as follows: initial denaturation at 95°C for 5 minutes, followed by 30-35 cycles of

1) denaturation at 94°C for 1 minutes, 2) annealing at 58°C for 30 seconds and 3) extension at

72°C for 30 seconds each. The final extension was at 72°C for 10 minutes. A matrix of forward

and reverse primer sets were designed on different exon hits and were used in separate reactions

89

to amplify the gene from the MTs cDNA library (Table 2). To search the MTs cDNA library for

regions upstream and downstream of the cloned internal sequence, gene-specific forward and

reverse primers was created over the internal sequence. The forward primer was used in PCR

with a vector-specific reverse primer which was generated over the plasmid that used to generate

the cDNA library: pDNR-LIB 3 -88 rev (AGTCATACCAGGATCTCCTAGGG). This primer is

88bp away from the restriction site of the vector. Subsequently, the product was purified and

nested PCR reactions were also done using nested reverse 2 and reverse 3 primers with pDNR-

LIB 3 -55 rev (GACCATGTTCACTTACCTACTGG) and pDNR-LIB 3 -25 rev

(GCCAAACGAATGGTCTAGAAAG) vector specific primers respectively, to amplify specific

3’ ends. Similarly, upstream regions of the gene were amplified using gene-specific reverse

primers and vector-specific forward primers: DNR-libfor1 (GTGGATAACCGTATTAC-

CGCC) and DNR-libfor2 (ACGGTACCGGACATATGCC). For all positive and negative

controls for PCR, ribosomal (rp49) primers rp49_qPCR-for (GTGAAACTCAGGAGAAATT

GGC) and rp49-rev2 (AGGACACACCATGCGCTATC) were used. Besides screening the

cDNA library, 5’ and 3’ Rapid Amplification of cDNA Ends (RACE) was also performed using

a 5’/3’ RACE kit (Roche Applied Science, Mannheim, Germany). All PCR samples were

visualized using ethidium bromide-stained 1% (w/v) agarose gels. Any bands of interest were gel

extracted or column purified using the EZ-10 Spin Column DNA Gel Extraction Kit (Bio Basic

Inc., Markham, ON, Canada) and EZ-10 Spin Column PCR Products Purification Kit (Bio Basic

Inc., Markham, ON, Canada) respectively. PCR extracts were cloned into the pGEM®-T vector

(Promega Corporation, Madison, WI, USA) and later transformed into competent Mach1™ T1R

Escherichia coli cells (Life Technologies Corporation, Carlsbad, CA, USA). Through ampicillin-

selective plating, transformed cells specifically containing the recombinant vector were picked

and grown into overnight cultures. Next, plasmid DNA was extracted from them and sequenced

at the SickKids DNA Sequencing Facility (The Centre for Applied Genomics, Hospital for Sick

Children, Toronto, ON, Canada).

90

RNA Isolation, cDNA synthesis & Reverse-Transcription PCR (RT-PCR)

Around 10 R. prolixus were dissected at each time and RNA from the tissues was

extracted using the SV Total RNA Isolation kit (Promega Corporation, Madison, WI, USA). For

the feeding experiments, the insects were fed on saline with ATP and were immediately

dissected. Subsequently, single stranded cDNA was synthesized using the Oligo d(T)20 primer

and approximately 400ng of RNA was used as template through the iScript™ Select cDNA

Synthesis kit (Bio-Rad Laboratories Ltd., Mississauga, ON, Canada). For the cDNA synthesis

reactions, the thermal cycler was programmed to prime at 25°C for 5 minutes, reverse

transcription at 42°C for 30 minutes and then inactivation of enzyme at 85°C for 5 minutes. A

1:2 dilution was done of the synthesized cDNA and an aliquot of this was subsequently used as

template in a PCR to test for the expression of the receptor gene in different tissues. The thermal

cycler conditions for the PCR were similar to the conditions programmed for regular PCR

mentioned above.

Bioinformatics

All the sequencing results were used to generate a Contig for the 5-HT receptor isolated from the

MTs cDNA library. Any discrepancies amongst the sequencing results were removed and noted

for future reference since they might be errors made by the non-proofreading Taq polymerase.

The Rhodnius preliminary genome database on Geneious was also searched (via tBLASTn)

using the consensus Contig sequence for comparison. This sequence was also inputted as a query

in protein BLAST on the ExPASy website to analyze its similarities with protein sequences from

other organisms. Exon-Intron splice sites were confirmed using online Berkeley splice site

prediction software (http://www.fruitfly.org/seq_tools/splice.html) (Reese, 2007). Multiple

sequence ClustalW alignments were created using a few of the highest scoring BLAST hits via

the online ClustalW2 software (http://www.ebi.ac.uk/Tools/clustalw2/index.html) (Larkin et al.,

2007). The BoxShade version 3.21 was used to illustrate amino acid conservation in the multiple

sequence alignment (http://www.ch.embnet.org/software/BOX_form.html). Also,

transmembrane domain (TMD) predictions were made using the TMHMM server version 2.0

(http://www.cbs.dtu.dk/services/TMHMM/) (Krogh et al., 2001; Sonnhammer et al., 2007).

91

Results

A Partial Nucleotide Sequence Encoding a Putative 5-HT Receptor: SK_Rhopr5HT2R

Initially, a Contig consensus sequence of 1594bp encoding a 516 amino acid ORF was retrieved

from Song Kim, Orchard Lab (Figure 1). This Contig sequence was previously created using

partial sequences obtained from 5’ and 3’ RACE. It had both a start and stop codon but lacked 5’

and 3’ untranslated regions (UTRs). The online transmembrane prediction tool (TMHMM)

predicted 7 TMDs which are displayed in figure 1. Protein BLAST analysis on the online

ExPASy software demonstrated that this sequence shares similarities with both 5-HT2 receptors

and dopamine receptors from some invertebrate and vertebrates, including a putative 5-HT2α

receptor from the honeybee A. mellifera (Ac: E5BBP1). tBLASTn analysis with the Rhodnius

preliminary genome database illustrated that the entire consensus sequence was situated in

Contig 4 of the genome database, with 5 potential intron regions spanning between 6 exons. The

pre-existing primers were used to amplify and isolate the entire SK_Rhopr5-HT2R cDNA (Table

1). However, this gene could not be isolated in its entirety and most primer combinations did not

amplify anything from the MTs cDNA library.

Figure 2 displays the primer combinations that did work and amplified segments of the gene, the

largest gene segment being a 900bp region amplified by the primers: 5HTR2 430-495F1and

Jpp5HTR2-R1 and the second largest region being a 400bp region that was amplified using

5HTR2 349-424F1 and 5HTR2 430-495R1 primers. Since these primer combinations gave the

two largest segments of the gene, they were used to design primers for the subsequent PCR

assays to screen the cDNA library. It is important to note that amplification of these specific gene

segments was not consistent in multiple trials and sometimes, very faint or no banding was

observed via gel electrophoresis. Moreover, both 5’ and 3’ RACE failed to amplify the UTRs of

this putative 5-HT receptor cDNA (data not shown).

To study the presence of the SK_Rhopr5-HT2R transcript in different tissues, RT-PCR was

performed. Both primer sets mentioned above were tested and gave similar results. Figure 3

92

Figure 1. The SK_Rhopr5HT2R Contig consensus nucleotide sequence of 1594bp and its 517

amino acid ORF. Sequencing results obtained from screening the MTs cDNA library and 5’ and

3’ RACE were used to generate the Contig. Predicted 7 TMDs are highlighted in black whereas

the start and stop codons are bolded.

94

Figure 2. A gel electrophoresis image of a PCR testing different primer combinations that

amplified products from the MT cDNA library. Each lane corresponds to the different primers

used: lane 1—5HTR2 430-495F1 and Jpp5HTR2-R1, lane 2—5HTR2 430-495F1 and 430R1,

lane 3—5HTR2 430-495F2 and 430R2, lane 4—5HTR2 430-495F1 and 430R2, lane 5—5HTR2

430-495F2 and 430R1, lane 6—5HTR2 349-424F1 and 5HTR2 430-495R1, lane 7—5HTR2

349-424F1 and 5HTR2 430-495R2, lane 8—5HTR2 349-424F2 and 5HTR2 430-495R1 and lane

9—5HTR2 349-424F2 and 5HTR2 430-495R2. Rp49 primers were used for the positive and

negative control in lanes +ve and -ve. No template was used for the negative control.

96

Figure 3. A gel electrophoresis image of the RT-PCR results. Each lane represents cDNA from a

different tissue used as template. While lanes 1 and 2 are not part of the PCR assay of interest,

lanes 3-13 are reactions where primers 5HTR2 349-424F1 and 5HTR2 430-495R1 were used.

Lanes 14- 24 are indicative of control reactions where Rp49 primers were used. The cDNA used

in the lanes are as follows: lanes 3 and 14—Whole MT cDNA, lanes 4 and 15—DV, trachea and

fat body, lanes 5 and 16—CNS, lanes 6 and 17—Foregut, lanes 7 and 18—hindgut, lanes 8 and

19—salivary glands, lanes 9 and 20—ovary, lanes 10 and 21—testies, lanes 11 and 22 posterior

midgut, lanes 12 and 23—anterior midgut and lanes 13 and 24—negative control for cDNA

synthesis.

98

Figure 4. A gel electrophoresis image of an RT-PCR (n=2 replicates). Primers 5HTR2 349-

424F1 and 5HTR2 430-495R1 were used for lanes 1-7 whereas RP49 primers were used in

reactions in lanes 8-14 as positive controls. Lanes 1-3 and 8-10 represent cDNA template that

was synthesized from unfed R. prolixus whereas in reactions represented by lanes 4-6 and 11-13,

cDNA templates were synthesized from RNA extracted from fed insects. For the experiment and

control with fed and unfed R. prolixus, the cDNA templates used follow the consecutive order

per lane: MTs (lanes 1, 4, 8, 11), anterior midgut (lanes 2, 5, 9, 12), salivary glands (lanes 3, 6,

10, 13) and no template (lanes 7, 14).

100

specifically illustrates the RT-PCR results using 5HTR2 349-424F1 and 5HTR2 430-495R1

primers. Expression was observed in MTs, salivary glands, ovaries and testes (Figure 3). Once

again, these results were inconsistent in independent replicates.

Additionally, to test if the expression of the transcript is altered upon feeding, RT-PCR was also

done using RNA extracts from tissues derived insects fed on saline with ATP and another group

of 10 that were unfed prior to dissection, using the primers: 5HTR2 349-424F1 and 5HTR2 430-

495R1 primers. In this experiment, gel electrophoresis revealed no specific banding in the MTs,

salivary glands and anterior midgut reactions of both, the fed and unfed groups (Figure 4). All

positive controls using Rp49 primers worked, however, faint bands were also observed in the

negative controls (Figures 2-4).

A Second Attempt—Isolation of a Partial Putative 5-HT2 Receptor cDNA: GB_Rhopr5-HT2R

Since the above SK_Rhopr5-HT2R cDNA could not be isolated or extended from the MTs

cDNA library, another attempt was made to isolate the 5-HT2 receptor gene from scratch. First,

the 5-HT2B receptor gene from D. melanogaster was retrieved (Schaelinger et al., 2007). This

gene was compared with the preliminary genome database of R. prolixus using tBLASTn

analysis and gave many high similarity hits on Contig 4. These hits from the genome database

were then used to make new primer designs over the R. prolixus genome. Of all the primer

combinations, only one pair amplified a strong band: 5htr2-51.1for1 and 5htr2-51.1rev1 (i.e. Rest

of the primers that did not amplify products, are not shown) (Table 2). Subsequently, the region

encoded between these primers was analyzed via protein BLAST on ExPASy, with a high

similarity to a protein sequence from D. mojavensis (Ac: B4KCQ3) that is similar to a 5-HT2

receptor from D. melanogaster but has not been annotated as such on ExPASy and NCBI. The

gene sequence from D. mojavensis was compared to the Rhodnius preliminary genome database

and once again, new primers were generated over the highest similarity hit regions of the genome

database (Table 2). A matrix of primer combinations was tested to amplify segments of varying

lengths of the gene from the MTs cDNA library (Figure 5). All primer combinations amplified

specific gene products except for the primers: SR287.3F1 - 5htr251.1rev1 and SR273.0F1 -

73.0R1. The primers SR2730.F1 and 5htr251.1rev1 primers amplified a large product from

101

Figure 5. A gel electrophoresis image of a PCR testing different primer combinations generated

over regions that are similar to a protein from D. mojavensis (Ac: B4KCQ3). The MT cDNA

library was used as template for each lane. The primers used in each reaction are as follows: lane

1—SR287.3F1 & 87.3R1, lane 2—SR287.3F1 & 5htr251.1rev1, lane 3—SR273.0F1 & 73.0R1,

lane 4—SR273.0F1 & 5htr251.1rev1, lane 5—SR260.0F1 & 60.0R1, lane 6—SR260.0F1 &

5htr251.1rev1, lane 7—SR258.0F1 & 58.0R1, lane 8—SR258.0F1 & 5htr251.1rev1, lane 9—

SR258.0F2 & 58.0R2 and lane 10—SR258.0F2 & 5htr251.1rev1. The lanes 11 and 12 are the

positive and negative controls, where Rp49 primers were used. No template was used for the

negative control.

103

Figure 6. Picture of a gel electrophoresis of an RT-PCR reaction. Different cDNA was used as

template for each lane. For the experimental reactions on the top lane, SR273.0F1 and

5hhtr251.1rev1 primers were used whereas for the control lane at the bottom, the Rp49 primers

were used once again. The cDNA used in each reaction is listed as follows: lane 1—CNS, lane

2—foregut, lane 3—salivary glands, lane 4—hindgut, lane 5—ovaries, lane 6—testis, lane 7—

DV, fat body and diaphragm, lane 8—MTs, lane 9—posterior midgut, lane 10—anterior midgut,

lane 11—no template.

105

the MTs cDNA library. Once this region was successfully sequenced, gene-specific primers

overlaying it were used with vector-specific primers to screen the cDNA library for the missing

upstream and downstream regions of the transcript (Table 2).

RT-PCR revealed a large gene-specific transcript (i.e. representing cDNA iv in Figure 8) is

expressed in the central nervous system (CNS), foregut, salivary glands and the MTs of R.

prolixus (Figure 6). In fact, gel electrophoresis displayed two closely situated bands in the lanes

representing the above cDNA. Similar double banding was also observed in 3’RACE results

(data not shown).

By sequencing amplified products of RACE and screening the cDNA library, a new Contig

consensus sequence was obtained. This sequence was 2245bp and encoded a single ORF of 741

amino acids (Figure 7). This Contig sequence encodes a stop codon at the 3’ end but it is still

missing a polyadenylation signal (i.e. AATAAA in the 3’UTR) at the 3’ end as well as a start

codon and a 5’ UTR at the 5’ end. The online TMHMM software again predicts 7 TMDs to be

encoded by ORF. So far, based on exon-intron boundaries and analysis of the partial gene, it is

structurally composed of 6 exons and 5 introns (Figure 8).The largest of the exons is the last

exon (6) while the first exon has only been sequenced partially. Most of the introns are extremely

large, approximating between 3.14kb to 44.3kb. The ORF also contains a highly conserved DRY

motif in the predicted 3rd intracellular loop of this predicted serotonin receptor, which is also

characteristic of a GPCR (Gether et al., 2002).

Attempts to amplify the entire GB_Rhopr5-HT2R using gene-specific primers were unsuccessful

and only overlapping segments of the Contig were amplified. However, two large overlapping

regions were isolated using primer combinations: 1) 5htr258.2for1 and SR260.0R1 and 2)

SR2730.0 and 5htr251.1rev1 which represent cDNAs i and iv respectively (Figure 8). No PCR

products were generated when primers SR287.3F1 and 5htr251.1rev1 were tested (Figure 5),

which would otherwise amplify a cDNA spanning exons 2 to 6. Primers 5htr258.2for1 (on exon

2) and 5htr251.1rev1 (on exon 6) also failed to work (data not shown).

Protein BLAST analysis of this Contig sequence on ExPASy showed similarities with 5-HT2

receptors from several invertebrates. The greatest similarity was found to be to an orphaned

106

GPCR from Tribolium castaneum (Ac: D6WIY4) and the second closest hit was a 5-HT2β

receptor from A. mellifera (Ac: E5BBP2). Type 2 serotonin receptors from several other

invertebrates also shared similarities with the GB_Rhopr5HT2R including D. melanogaster (Ac:

Q8INQ6) and the crayfish, Procambarus clarkii (Ac: B0F4P8). Similarities were also observed

with vertebrate 5-HT2 receptors from species like the frog, Xenopus laevis, the mouse, Mus

musculus and humans (data not shown). Additional top hits were uncharacterized GPCR-like

proteins from different species of Drosophila.

A multiple sequence alignment generated via the online ClustalW software displays the amino

acid conservation of the 5-HT receptors of the invertebrates mentioned above and the ORF of the

putative 5-HT receptor gene (GB_Rhopr5HT2R) (Figure 9). Highly conserved regions are

observed specifically in regions predicted as TMDs.

107

Figure 7. GB_Rhopr5HT2R Contig consensus nucleotide sequence of 2245bp and a potential

ORF comprising of 741 amino acids, based on all the 5’and 3’ RACE products and internal

sequences that have been sequenced thus far. The 7 TMDs have been highlighted in black

whereas the stop codon is bolded.

109

Figure 8. Schematic depicting the genetic structure of the GB_Rhopr5-HT2R Contig which

encodes for a putative 5-HT2 receptor. Exons are boxed (numbered in circles), introns are

displayed as lines connecting the boxes while the 5’ and 3’ UTRs are displayed as dash lines.

The sizes of exons are displayed within the boxes and the sizes of introns are displayed below the

introns marked by dashed arrows. Finally, the red arrows (numbered on the left) located above

the schematic represent segments of cDNA (i.e. excluding the introns) that were amplified and

used to generate the Contig. This schematic is not drawn to scale.

111

Figure 9. ClustalW2 alignment of the GB_Rhopr5HT2R Contig sequence obtained via cloning

along with 5-HT2 receptor sequences from D. melanogaster sequence (Ac: Q8INQ6), A.

mellifera (Ac: E5BBP2), P. clarkii (Ac: B0F4P8) as well as an uncharacterized GPCR from T.

castaneum (Ac: D6WIY4). The 7 TMDs are boxed with a number indicated above. Identical

amino acid residues are highlighted in black whereas similar amino acids in a column are

highlighted in grey. Only similarities and/or identities that occur in 50% of the cases in each

column are indicated. The highly conserved DRY motif located right after the 3rd TMD is

underlined.

113

Discussion

The objective of this study was to isolate and characterize a 5-HT receptor from the MTs of R.

prolixus as well as to study its expression in different tissues since 5-HT is involved in

modulating post-prandial diuresis and feeding activities of R. prolixus. Initially, a 1594bp Contig

consensus sequence (i.e. SK_Rhopr5HT2R) encoding a 516 amino acid ORF, was predicted as a

potential 5-HT receptor transcript (Figure 1). Protein BLAST analyses showed high similarities

of this sequence with 5-HT type 2 receptors from different organisms like the honeybee A.

mellifera (Ac: E5BBP1) (Figure 1). However, failure to isolate the entire gene transcript using

gene-specific primers as well as the inconsistencies in amplifying gene-specific products from

different tissue-derived cDNA in the RT-PCR experiments, ruled out the possibility of this

Contig sequence being the 5-HT2 receptor involved in rapid diuresis.

Nevertheless, before abandoning this Contig sequence, another hypothesis was developed. It was

predicted that the faint banding observed in PCR and RT-PCR gels was attributed to low

transcript levels of the receptor in non-fed insects. Since the cDNA library and individually

isolated cDNA were synthesized from RNA of non-fed insects, it was postulated that perhaps the

5-HT receptor transcript is up-regulated in tissues during feeding due to the receptor’s significant

role in diuresis and digestion. However, this hypothesis was inconclusive since the gene could

not be amplified in both the fed and unfed groups (Figure 4). During the experiments, it was also

noticed that the negative controls had faint bands in them (Figures 2-4). However, this was due

to contamination of the primer working and master stocks. To avoid this, smaller aliquots of

primer working stocks were made and filter pipette tips were used cautiously, avoiding

contamination. Nonetheless, more independent replicates need to be repeated to confirm these

observations. It is also plausible that this receptor type is not highly expressed in 5th instar R.

prolixus and could in fact be involved in the physiology pertaining to other developmental stages

of this insect. For example, the 5-HT2 receptor is highly expressed during embryogenesis,

specifically gastrulation of D. melanogaster (Colas et al. 1995; see Nebigil et al., 2001).

Likewise, it is possible that this receptor subtype is involved in the embryonic stage of

development or even in adulthood. Since we are specifically trying to identify the 5-HT receptor

involved in the dramatic period of diuresis and digestion of 5th instars, the SK_Rhopr5HT2R

114

Contig was set aside for later studies and the search for the receptor of interest was initiated

again, from scratch.

So, in the second part of this study, new primer designs were created using D. melanogaster 5-

HT2B receptor and a D. mojavensis sequence. Sequencing results were used to generate a new

Contig sequence that was 2245bp and encoded a single ORF of 741 amino acids, including a stop

codon (Figure 7). Topology predictions indicate that the deduced protein encodes for 7 TMDs

and small segments of the extracellular N-terminus and the intracellular C-terminus (data not

shown). It is worth noting that this gene is also located on Contig 4 of the genomic database and

is downstream of SK_Rhopr5HT2R. In fact, there are many regions on this Contig that share

similarities with 5-HT receptors from different insect species as well as the crustacean, P. clarkii

(not shown). This could suggest that genes encoding different 5-HT receptor types are located in

close proximity of each other, possibly on the same chromosome. Unlike the ORF of

SK_Rhopr5HT2R, the protein encoded by GB_Rhopr5HT2R contains a highly conserved DRY

motif predicted in the 3rd intracellular loop of the GPCR. This D/E RY motif is characteristic of

GPCRs as it is involved in an ionic lock which is broken during receptor activation (Gether et al.,

2002). This further supports that GB_Rhopr5HT2R does in fact encode for a GPCR; whereas, the

absence of this motif in the ORF encoded by SK_Rhopr5HT2R, raises speculation on whether

this Contig does encode for a GPCR or not. Nevertheless, the knowledge of GPCRs is largely

based on vertebrate studies and thus, more research on invertebrate GPCRs is required to confirm

this.

Strong and specific bands were amplified from cDNA of MTs, CNS, foregut and salivary glands

in the RT-PCR experiments, therefore indicating the presence of the transcript in these tissues

(Figure 6). Serotonin is known to target all of these tissues and thus, the presence of the

transcript in these tissues is justifiable (see Orchard et al., 2006; 2009). Serotonin is also known

to target the integument, hindgut and dorsal vessel but the transcript was absent in these tissues.

The possibility of tissue-specific serotonin receptor (sub)type expression could account for this

since these families of receptors work through different signal transduction mechanisms (see

Raymond et al., 2001; Tierney, 2001). There was also no amplification of a product in the

anterior midgut, although previous research indicates that the 5-HT2 receptor is most likely found

in the anterior midgut. Levels of cAMP elevate upon application of serotonin on 5th instar

115

anterior midgut (Te Brugge et al., 2009) as well as those of adults (Barret et al., 1993). In adults,

serotonin is also involved in the up-regulation of a gene involved in lipid metabolism (Alves-

Bezerra et al., 2010). Possibly, a different receptor type (e.g. type 1 or 7) is involved in fluid

absorption across the anterior midgut of 5th instars. Protein BLAST analyses on ExPASy shows

similarities between the predicted ORF and 5-HT2 receptors from different invertebrates and

vertebrates therefore elucidating the function of this protein as a serotonin receptor. Moreover,

previous pharmacological studies using mammalian 5-HT type 2 receptor antagonists like

ketanserin, mianserin and spiperone indicate that the likelihood of finding the 5-HT2 receptor in

the MTs, midgut and integument is the greatest (Barret et al., 1993; Maddrell et al., 1991).

Similar pharmacological studies have also predicted the presence of the 5-HT2 receptor in the

locust midgut (Molaei and Lange, 2003) and cockroach salivary glands (Troppmann et al., 2007).

Research on signal transduction pathways of these receptors also propose that the 5-HT receptor

found in the MTs and midgut most likely follows a cAMP-dependent second messenger pathway

because serotonin causes a dose-dependent rise in cAMP and simultaneously increases the rate of

secretion in the tubules, hence supporting our predictions (Barret et al., 1993; Te Brugge et al.,

2009;). Although the signal transduction pathway of this receptor has yet to be confirmed, it is

important to note that the characterization of these 5-HT receptors in insects; as types 1, 2 and 7,

are greatly dependent on pharmacological studies using vertebrate agonists and antagonists.

There is little knowledge on pharmacological characteristics of insect 5-HT receptors and there

could be many differences between the vertebrate and invertebrate receptors that are not

accounted for in these studies. For instance, the 5-HT2 receptor in insects has been predicted to

follow the cAMP-dependent pathway whereas in vertebrates, it follows a phospholipase C

pathway (see Raymond et al.; Tierney, 2001). Accordingly, more in-depth studies on 5-HT

receptors in invertebrates are required to enhance pharmacological knowledge pertaining to

invertebrates and will also provide a more concise and accurate standard nomenclature for

insects.

The inability to amplify the entire GB_Rhopr5-HT2R cDNA could have several reasons. The

inefficiency of the primers could be possible especially since this gene is very large (Figure 8).

Another reason could be the possibility of alternative splicing. Gel Electrophoresis of RACE and

RT-PCR reactions revealed the presence of two bands, with the use of certain primer sets (Figure

116

6). This could indicate the presence of multiple transcript variants of this 5-HT2 receptor-like

transcript as a result of alternative splicing. Thus, it is possible that the reason the entire cDNA

could not be amplified is due to primers created over a spliced intron regions. The alternatively

spliced variants and their translated isoforms may be expressed, distributed and/or utilized

differentially. D. melanogaster for example, has 5 different isoforms of the 5-HT2 receptor (Ac:

Q9VN38.1).

The ClustalW alignment also reaffirms the likelihood that the new Contig sequence is most

likely a 5-HT2 receptor, because it shares similarities with 5-HT2 receptors from different

species. The ClustalW alignment of the ORF of the new Contig sequence with 5-HT2 receptors

from D. melanogaster gene, A. mellifera and P.clarkii illustrates that these proteins share a high

degree of conservation in amino acids, particularly in the regions predicted as the 7 TMDs

(Figures 9). The greatest similarity of the ORF of GB_Rhopr5HT2R is to an uncharacterized

protein from T. castaneum, which is most likely a GPCR that has yet to be deorphanized as a

serotonin receptor. In addition, the greatest variability with the multiple sequence alignment is

observed between TMD 5 and 6. It is difficult to predict how much of the protein is still missing

at the N-terminus because these proteins vary in size. In fact, the ORF encoded by the

GB_Rhopr5HT2R Contig also shares similarities with truncated 5-HT2 receptors: a 383 amino

acid protein from A. aegypti (Q1DGP1) and a 416 amino acid protein from D. melanogaster

(Q9VHP6), which encode for 2 and 5 TMDs respectively (not shown). This further demonstrates

the possibility of alternative splicing of this gene.

Despite many attempts, the whole cDNA of the GB_Rhopr5HT2R Contig could not be amplified,

most likely due to inefficient primers and/or alternative splicing. However, unlike the

SK_Rhopr5HT2R Contig, sequencing of the overlapping segments as well as RT-PCR of the

GB_Rhopr5HT2R Contig gave consistent results. Thus, molecular cloning, spatial expression

studies and bioinformatics analysis also illustrate promising results which suggest that the

GB_Rhopr5HT2R Contig does in fact encode for a 5-HT2 receptor in 5th instar R. prolixus. This

receptor may play a crucial role in the rapid diuresis which is associated to the disease

transmission and thus, further experimentation will be essential in determining its potential as a

pharmacological target in disease prevention.

117

Future Directions

The most immediate step in this project will be to generate efficient primers over exon-exon

boundaries in order to identify splice variants because the inability to amplify the whole receptor

cDNA and the double banding observed in RT-PCR and RACE hint on the possibility of

alternative splicing. The remaining 5’ and 3’ ends of the gene will be retrieved via RACE, in

order to isolate the entire gene. Northern blots will also provide a better estimation of the size of

the transcript. Expression levels and localization of the gene will be studied via quantitative PCR

(qPCR), florescent in situ hybridization whereas localization of the protein can be confirmed via

immunohistochemistry. Repeating the RT-PCR will reconfirm the gene expression in different

tissues. Functional assays performed by transfecting Chinese hamster ovarian cells with this gene

will further enhance our knowledge of this receptor and its specificity for serotonin, its agonists,

antagonists and other chemicals and neuropeptides.

To gain a better understanding of the biochemistry behind the 5-HT receptor in R. prolixus and

other insects, the structure and receptor’s intracellular pathways are also areas of research to look

into in future research. Furthermore, immunohistochemistry in conjuction with qPCR can also

provide insight on time course expression during the process of feeding. These studies can

provide more comprehensive details on the differences between vertebrate and invertebrate 5-HT

receptors and can also improve the pharmacological studies of 5-HT receptors in insects. In

conclusion, in congruence with results of the current research, further studies will enhance the

understanding of the role of the serotonin receptor in regulating the digestive and feeding

activities of the hemipteran insect, R. prolixus.

118

References

Ali, D.W., Orchard, I. and Lange, A.B. (1993). The aminergic control of locust (Locusta

migratoria) salivary glands: evidence for dopaminergic and serotonergic innervation. Journal

of Insect Physiology. 8, 623-632.

Alves-Bezerra, Majerowicz, D., Grillo, L.A.M., Tremonte, H., Almeida, C.B., Braz, G.R.C.,

Sola-Penna, M., Paiva-Silva, G.O., and Gondim, K.C. (2010). Serotonin regulates an acyl-

CoA-binding protein (ACBP) gene expression in the midgut of Rhodnius prolixus. Insect

Biochemistry and Molecular Biology. 40, 119-125.

Barret, F.M., Orchard, I and TeBrugge, V. (1993). Characteristics of serotonin-induced cyclic

AMP elevation in the integument and anterior midgut of the blood-feeding bug, Rhodnius

prolixus. Journal of Insect Physiology. 39, 581-587.

Blenau, W. and Thamm, M. (2011). Distribution of serotonin (5-HT) and its receptors in the

insect brain with focus on the mushroom bodies. Lessons from Drosophila melanogaster and

Apis mellifera. Arthropod Structure & Development. 40, 381-394.

Coast, G. (2007). The endocrine control of salt balance in insects. General and Comparative

Endocrinology. 152, 332-338.

Colas, J.F., Launay, J.M. and Kellermann, O. (1995). Drosophila 5-HT2 serotonin receptor:

coexpression with fushi-tarazu during segmentation. Proceedings of the National Academy of

Sciences of the United States of America. 92, 5441-5445.

Dacks, A.M., Dacks, J.B., Christensen, T.A. and Nighorn, A.J. (2006). The cloning of one

putative octopamine receptor and two serotonin receptors from the tobacco hawkmoth,

Manduca sexta. Insect Biochemistry and Molecular Biology. 36, 741-747.

Falibene, A., Rössler, W. and Josens, R. (2012). Serotonin depresses feeding behaviour in ants.

Journal of Insect Physiology. 58, 7-17.

119

Gether, U., Asmar, F., Meinild, A.K. and Rasmussen, S.G.F. (2002). Structural basis for

activation of G-protein-coupled receptors. Pharmacology & Toxicology. 91, 304-312.

Heslop, J.P., and Berridge, M.J. (1980). Changes in cyclic AMP and cyclic GMP

concentrations during the action of 5-hydroxytryptamine on an insect salivary gland.

Biochemical Journal. 192, 247-255.

Johnson, O., Becnel, J. and Nichols, C.D. (2011). Serotonin receptor activity is necessary for

olfactory learning and memory in Drosophila melanogaster. Neuroscience. 192, 372-381.

Krogh, A.,Larsson, B., Heijne, G. and Sonnhammer, L. (2001). Predicting transmembrane

topology with a hidden Markov model: Applicaion to complete genomes. Journal of

Molecular Biology. 3053, 567-580.

Lange, A.B. Orchard, I. and Barrett, F.M. (1989). Changes in haemolymph serotonin levels

associated with feeding in the blood-sucking bug, Rhodnius prolixus. Journal of Insect

Physiology. 35, 393-399.

Larkin, M.A., Blackshields G., Brown N.P., Chenna R., McGettigan P.A., McWilliam H.,

Valentin F., Wallace I.M., Wilm A., Lopez R., Thompson J.D., Gibson T.J. and Higgins

D.G. (2007) ClustalW and ClustalX Version 2. Bioinformatics. 23, 2947-2948.

Lent, C.M. and Dickinson, M.H. (1988). The neurobiology of feeding in leeches. Scientific

American. 258, 98-103.

Lima, F.M., Oliveira, P., Mortara, R.A., Silveira, J.F. and Bahia, D. (2010). The challenge of

Chagas’ disease: Has the human pathogen, Trypanosoma cruzi, learned how to modulate

signalling events to subvert host cells? New Biotechology. 27, 837-843.

Maddrell, S.H.P., Herman, W.S., Mooney, R.L. and Overton, J.A. (1991). 5-

Hydroxytryptamine: a second diuretic hormone in Rhodnius prolixus. Journal of

Experimental Biology. 156, 557-566.

120

Martini, S.V., Nascimento, S.B. and Morales, M.M. (2007). Rhodnius prolixus Malpighian

Tubules and control of diuresis by neurohormones. Annals of Brazilian Academy of Sciences.

79, 87-95.

Molaei, G. and Lange, A.B. (2003). The association of serotonin with the alimentary canal of

the African migratory locust, Locusta migratoria: distribution, physiology and

pharmacological profile. Journal of Insect Physiology. 49, 1073-1082.

Nässel, D.R. (1988). Serotonin and serotonin-immunoreactive neurons in the nervous system of

insects. Progress in Neurobiology. 30, 1-85.

Nebigil, C.G., Etienne, N., Schaerlinger, B., Hickel, P., Launay, J.-M., and Maroteaux, L.

(2001). Developmentally regulated serotonin 5-HT2B receptors. International Journal of

Developmental Neuroscience. 19, 365-372.

O’Donnell, M.J. (2009). Too much of a good thing: how insects cope with excess ions or toxins

in the diet. The Journal of Experimental Biology. 212, 363-372.

O’Donnell, M.J. and Spring, J.H. (2000). Modes of control of insect Malpighian tubules:

synergism, antagonism, cooperation and autonomous regulation. Journal of Insect Physiology.

46, 107-117.

Orchard, I., Lange, A.B. and Barrett, F.M. (1988). Serotonergic supply to the epidermis of R.

prolixus: evidence for serotonin as the plasticising factor. Journal of Insect Physiology. 34,

873-879.

Orchard, I., Lange A.B., Cook, H. and Ramirez, J.M. (1989). A subpopulation of dorsal

unpaired median neurons in the blood-feeding insect, Rhodnius prolixus, displays serotonin-

like immunoreactivity. Journal of Comparative Neurology. (289), 118-128.

Orchard, I. (2006). Serotonin: a coordinator of feeding-related physiological events in the

blood-gorging bug, Rhodnius prolixus. Comparative Biochemistry and Physiology. 144, 316-

324.

121

Paluzzi, J.P., Naikkhwah, W. and O’Donnell, M.J. (2012). Natriuresis and diuretic hormone

synergism in R. prolixus upper Malpighian tubules is inhibited by anti-diuretic hormone

Rhopr-CAPA-α2. Journal of Insect Physiology. 58, 534-542.

Paluzzi, J.P., Russell, W.K., Nachman, J. and Orchard, I. (2008). Isolation, cloning and

expression mapping of a gene encoding an antidiuretic hormone and other CAPA-related

peptides in the disease vector, Rhodnius prolixus. Endocrinology. 149, 4638-4646.

Raymond, J.R., Mukhin, Y.V., Gelasco, A., Turner, J. Collinsworth, G., Gettys, T.W.,

Grewal, J.S. and Garnovskaya, M.N. (2001). Multiplicity of mechanisms of serotonin

receptor signal transduction. Pharmacology & Therapeutics. 95, 179-212.

Reese M.G., Eeckman, F.H., Kulp, D. and Haussler, D. (1997). Improved splice site

detection in genie. Journal of Comparative Biology. 4, 311-323.

Schaerlinger, B, Launay, JM, Vonesch, JI, and Maroteaux, L. (2007). Gain of affinity point

mutation in the serotonin receptor gene 5-HT2Dro accelerates germband extension

movements during Drosophila gastrulation. Developmental Dynamics. 236, 991-999.

Sonnhammer, ELL., Heijne, G. and Krogh, A. (1998). A hidden Markov model for

predicting transmembrane helices in protein sequences. Proceedings of the Sixth

International Conference on Intelligent Systems for Molecular Biology: AAAi Press: Menlo

Park, CA, pages 175-182.

Te Brugge, V.A., Lombardi, V.C., Schooley, D.A. and Orchard, I. (2005). Presence and

activity of a Dippu-DH31-like peptide in the blood-feeding bug, Rhodnius prolixus. Peptides.

26, 29-42.

Thamm, M., Balfanz, S., Scheiner, R., Baumann, A. and Blenau, W. (2010).

Characterization of the 5-HT1A receptor of the honeybee (Apis mellifera) and involvement

of serotonin in phototactic behaviour. Cellular and Molecular Life Sciences. 67, 2467-2479.

122

Thompson, K.S.J., Rayne, R.C., Gibbon, C.R., May, S.T., Patel, M., Coast, G.M. and

Bacon, J.P. (1995). Cellular colocalization of diuretic peptides in locusts: a potent control

mechanism. Peptides. 16, 95–104.

Tierney, A.J. (2001). Structure and function of invertebrate 5-HT receptors: a review.

Comparative Biochemistry and Physiology Part A. 128, 791-804.

Troppmann, B., Blafanz, S., Baumann, A. and Blenau, W. (2010). Inverse agonist and

neutral antagonist actions of synthetic compounds at an insect 5-HT1 receptor. Bristish

Journal of Pharmacology. 159, 1450-1462.

Troppmann, B., Walz, B. and Blenau, W. (2007). Pharmacology of serotonin-induced

salivary secretion in Periplaneta americana. Journal of Insect Physiology. 53, 774-781.

von Nickisch-Rosenegk, E., Krieger, J., Kubick, S., Laage, R., Strobel, J., Strotmann, J.

and Breer, H. (1996). Cloning of biogenic amine receptors from moths (Bombyx mori and

Heliothis virescens). Insect Biochemistry and Molecular Biology. 26, 817-827.

World Health Organization (WHO). (2008). Chagas disease: control and elimination—Report

of the Secretariat. Executive Board, Session 124th. Provisional agenda item 4.14, 1-4.

the physiological roles of rhopr-kinins and the molecular ... · garima bhatt master of science...

Documents