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Neuropeptide Mimetics: The Physiological Effects of Kinin and CAPA Analogs in Rhodnius prolixus
by
Vishal Sangha
A thesis submitted in conformity with the requirements for the degree of Master of Science
Cell and Systems Biology University of Toronto
© Copyright by Vishal Sangha 2019
ii
Neuropeptide Mimetics: The Physiological Effects of Kinin and CAPA
Analogs in Rhodnius prolixus
Vishal Sangha
Master of Science
Cell and Systems Biology University of Toronto
2019
Abstract
In the Chagas disease vector Rhodnius prolixus, the kinin and CAPA neuropeptides
modulate a host of feeding and diuresis-related behaviours that are implicated in disease
transmission. CAPA and kinin neuropeptide analogs have been developed to elicit potent changes
in physiology, to be later incorporated in novel pest-control strategies. Here, the effects of kinin
and CAPA analogs were investigated on feeding and diuresis-related tissues, with the kinin and
CAPA analogs inducing physiological changes in vivo and in vitro. Within the hindgut, novel
intracellular interactions were uncovered between RhoprCAPA, Rhopr-kinin, and serotonin [5-
hydroxytryptamine (5-HT)]. Following identification and sequencing of the Rhopr-kinin receptor,
the receptor transcript was observed throughout the gut, with RNA interference (RNAi)-mediated
knockdown of the receptor causing reductions in hindgut contractions and increasing the size of
blood meal consumed. Overall, these findings highlight the role of kinin and CAPA within R.
prolixus, and the promise of their neuropeptide analogs to be used as lead compounds in pest-
control strategies.
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Acknowledgments
I am truly grateful to Dr. Lange for providing me with the opportunity to conduct research in the
esteemed Lange/Orchard lab. Your guidance and endless support throughout these past two years
has allowed me to grow as a researcher, and as a person. To Dr. Orchard, who essentially served
as a co-supervisor, thank you for taking the time out of your busy schedule to provide direction
and feedback on my work, and answering every possible silly question of mine. Yours and Dr.
Lange’s passion for science is truly inspiring.
I would like to thank Dr. Senatore for being a part of my committee and providing me with
feedback throughout my research, and Dr. Westwood for serving as an external examiner
To past and present members of the Lange/Orchard lab who I have had a chance to work with,
thank you for welcoming me with open arms and being wonderful colleagues. We have created
some delightful memories, and I am glad to know that I am leaving this lab with some great friends.
Lastly, I’d like to thank my family for always supporting me regardless of what I decided to pursue.
To my parents, I am grateful for your encouragement and love, and emphasizing the value of hard
work and perseverance from a young age. To my brother Deepak, thank you for being a wonderful
role model, and a great brother. To Jaskaran, my best friend and partner in crime, thank you for all
of your love and support, and always believing in me even when I doubted myself. I am thankful
for all of you.
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Table of Contents Abstract…………………………………………………………………………….…………...ii
Acknowledgments…………………………………………………………………………...…iii
Organization of Thesis…………………………………………………………....…………....vi
List of Figures and Tables…………….……………………………………….…...…………vii
List of Abbreviations……………………………………….………………………...…….......ix
Chapter 1: General Introduction……………………………………………………......1
Rhodnius prolixus…………………………………………………………………………..........1
Neuropeptides……………………………………………………………………………….…...2
G Protein-Coupled Receptors (GPCRs) ……………………………………………….….….....4
Kinin……………………………………………………………………………………………..7
CAPA……………………………………………………………………………………...…....11
Post-Feeding Physiology………………………………………………………………...……..13
Diuresis…………………………………………………………………………….…...13
Excretion………………………………....………………………………………….….15
Neuropeptide Analogs………………………………………………………………………….16
Objectives of Thesis……………………………………………………………………………18
References………………………………………………………………………………….…..20
Chapter 2: Physiological Effects of Kinin and CAPA Analogs in the Chagas Disease Vector, Rhodnius Prolixus…………………………………………………….29
Abstract………………………………………………………………………………………...30
Introduction ……………………………………………………………………………………31
Materials and Methods……………………………………………………………………........34
Results ……………………………………………………………………………………........37
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Discussion……………………………………………………………………………….……..60
References…………………………………………………………………………….….…….66
Chapter 3: Identification and Cloning of the Kinin Receptor in the Chagas Disease Vector, Rhodnius Prolixus…………………………………………………….71
Abstract………………………………………………………………………………………...72
Introduction ……………………………………………………………………………………73
Materials and Methods……………………………………………………………...………….77
Results.………………………………………………………………………………………....82
Discussion………………………………………………………………….….……………….98
References……………………………………………………………………………………..104
Chapter 4: General Discussion………………………………………….……………..113
Feeding………………………………………………………………………………………...113
Kinin …………………………………………………………………………………..113
CAPA……………………………...…………………………………………………...115
Myotropic Effects……………………………………………………………………………...115
Kinin…………………………………………………………………………………...115
CAPA....…………………………………….………………………………………….117
Diuretic Effects…………………………………………………...……………………………126
Summary of Physiological Effects of Analogs…………...……………………………………129
References…………………………………………………………………………………...…131
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Organization of Thesis
This thesis consists of four chapters. Chapter 1 is a general introduction, providing background
information for the proceeding chapters. Chapter 2 is organized as a research article examining
the physiological effects of kinin and CAPA analogs and has been submitted to Insect
Biochemistry and Molecular Biology. Chapter 3 is also organized as a research article focusing
on the cloning of the R. prolixus kinin receptor, examining its expression profile, and RNAi-
mediated knockdown of the receptor transcript. Chapter 4 is a general discussion that discusses
the research findings of this thesis, and future directions.
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List of Figures and Tables
Chapter 1: General Introduction
Table 1: Amino acid sequences of Rhopr-kinins……………………………………………..…9
Chapter 2: Physiological Effects of Kinin and CAPA Analogs in the Chagas Disease Vector, Rhodnius Prolixus
Table 1: Structure of kinin and CAPA analogs………………………………………………...41
Figure 1: Effects of injection of Rhopr-kinin 2 and kinin analog on in vivo feeding and diuresis………………………………………………………………………………...……….42
Figure 2: Effects of injection of Rhopr-kinin 2 and CAPA analog on in vivo feeding and diuresis…………………………………………………………………………………………44
Figure 3: Effects of Rhopr-kinin 2 and kinin analog on hindgut basal tonus……………...….46
Figure 4: Effects of RhoprCAPA-2 and CAPA analog on hindgut basal tonus…………...….48
Figure 5: Potentiation effects of RhoprCAPA-2 and Rhopr-kinin 2 on hindgut basal tonus…50
Figure 6: Effects of co-application of Rhopr-kinin 2 and CAPA analog on the hindgut …….52
Figure 7: Potentiation effects of 5-HT and CAPA analog on the frequency of hindgut contractions…………………………………………………………………………………….54
Figure 8: Effects of co-application of 5-HT and RhoprCAPA-2 on the hindgut………….…..56
Figure 9: Effects of CAPA analog on Malpighian tubule secretion…………………………..58
Chapter 3: Identification and Cloning of the Kinin Receptor in the Chagas Disease Vector, Rhodnius Prolixus
Figure 1: RhoprKR cDNA sequence and exon map…………………………………………..86
Figure 2: Multiple sequence alignment of invertebrate kinin receptors and R. prolixus receptors……………………………………………………………….………………….........88
Table 1: Identity within transmembrane domains of invertebrate kinin receptors…………….91
Figure 3: Expression profile of the RhoprKR transcript…………………………………........92
Figure 4: Effects of RNAi-mediated RhoprKR knockdown on hindgut contractions………...94
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Figure 5: Effects of RNAi-mediated RhoprKR knockdown on in vivo feeding and diuresis……………………………………………………………………………………...….96
Supplemental Figure 1: Knockdown efficiency of RhoprKR in dsRhoprKR R. prolixus insects…………………………………………………………………………………………109
Table S1: Primers used for amplification of RhoprKR cDNA fragments…………………………………………………………………………….…….….109
Table S2: 5’ and 3’ RACE primers for RhoprKR……………………………………………110
Table S3: 5’ and 3’ primers used for dsRNA synthesis………………………………....…...110
Table S4: Primers used for qPCR analysis of the RhoprKR transcript……………………....111
Chapter 4: General Discussion
Figure 1: Model of kinin, CAPA, and 5-HT receptor activation……………………………..120
Figure 2: Model showing the intracellular interactions between RhoprCAPA-2 and Rhopr-kinin 2/5-HT…………………………………………………………………………………………122
Figure 3: Model showing the intracellular interactions between CAPA analog and Rhopr-kinin 2/5-HT……………………………………………………………………….………………...124
Figure 4: Summary of physiological effects of kinin and CAPA in the alimentary canal of R. prolixus………………………………………………………………………………………...127
Table 1: Summary of future directions………………………………………………...……...130
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List of Abbreviations 5-HT: Serotonin (5-hydroxytryptamine)
AC: Adenyl cyclase
ACE: Angiotensin converting enzyme
ACN: Acetonitrile
Aib: Alpha-aminoisobutyric acid
AMG: Anterior midgut
ANOVA: Analysis of variance
ARG: Ampicillin resistance gene
ATP: Adenosine triphosphate
BLAST: Basic local alignment search tool
Ca2+: Calcium
CaCl2: Calcium chloride
cAMP: Cyclic adenosine monophosphate
CAP: Cardioactive peptide
CCAP: Crustacean cardioactive peptide
cDNA: Complementary DNA
cGMP: Cyclic guanosine monophosphate
CHS: Chitin synthase
CNS: Central nervous system (CNS),
CRF/DH: Corticotropin-releasing factor-like diuretic hormone
CRZ: Corazonin
DH31: Calcitonin-like diuretic hormone
dsRNA: Double-stranded RNA
DV: Dorsal vessel
EC: Extracellular loop
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ER: Endoplasmic reticulum
FB: Fat body
FG: Foregut
GDP: Guanosine diphosphate
GPCR: G protein-coupled receptor
GTP: Guanosine triphosphate
HEK293/CNG: Human embryonic kidney cells stably expressing a cyclic nucleotide gated channel
HG: Hindgut
HPLC: High-performance liquid chromatography
IC: Intracellular Loop
IP3: Inositol triphosphate
KCl: Potassium chloride
MgCl2: Magnesium chloride
MT: Malpighian tubules
MTGM: Mesothoracic ganglionic mass
NaCl Sodium choloride
NaHCO3: Sodium bicarbonate
NEP: Neprilysin
ORF: Open reading frame
PBS: Phosphate-buffered saline
PIP2: Phosphatidylinositol 4,5-bisphosphate
PK: Pyrokinin
PLC: Phospholipase C
PMG: Posterior midgut
PVK: Periviscerokinin
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qPCR: Quantitative polymerase chain reaction
RACE: Rapid amplification of CDNA ends
RNAi: RNA Interference
SEM: Standard error of measure
SG: Salivary glands
SOG: Suboesophageal ganglion
TFA: Trifluoroacetic acid
TM: Transmembrane domains
UTR: Untranslated region
1
Chapter 1 General Introduction
Rhodnius prolixus
Rhodnius prolixus is a blood-gorging hemipteran of the Reduviidae family, domestic to
Central and South America (Dujardin et al., 1998; Buxton, 1930). Within the R. prolixus life cycle,
five nymphal stages (or instars) and a final adult stage exist, with the blood meal serving as a
requirement for successful moulting to occur between each stage and for reproductive processes
(Wigglesworth, 1934). This requirement of a blood meal to initiate developmental and
reproductive processes made R. prolixus an essential model insect in the very origins of insect
physiology and endocrinology (Davey, 2007). R. prolixus only requires one blood meal for the
transition through each life stage, surviving for months between feeds (Uribe, 1926; Wigglesworth,
1934). The transition period between each stage is predictable, during which essential
physiological processes of growth, development, ecdysis and reproduction can be thoroughly
examined (Azambuja et al., 2017). Native R. prolixus can be divided into two populations:
domiliciary, and sylvan. Sylvan populations tend to exist within palm tree leaves and pteridophytes
and are often found hidden with various species such as mammals, marsupials, and reptiles (Davey,
2007). The domiciliary population is more closely associated with humans, existing in damp, dark
spaces within houses, and tend to feed at night on humans or domestic animals (Davey, 2007;
Garcia et al., 2007). During feeding, R. prolixus consumes a blood meal that can be up to 8-10
times it’s body weight. This feeding strategy requires a tight regulation of osmotic balance, which
is subject to control by the neuroendocrine system (Orchard, 2006; Coast et al., 2002).
From a medical perspective, R. prolixus is an organism of interest, as it is a carrier of the
Trypanosoma cruzi parasite, making it a vector of Chagas disease (Dujardin et al., 1998; Moncayo,
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2003). Rhodnius prolixus is one of 12 Triatominae species that act as a vector for the T. cruzi
parasite (Schofield, 1988). Chagas disease, originally discovered by the Brazilian physician Carlos
Chagas, is principally found in the continental part of Latin America but has been recently detected
in the U.S, and Canada (WHO, 2019; Steverding, 2014). Six to seven million people worldwide
are infected with the T. cruzi parasite, with most cases in Latin America (WHO, 2019). Chagas
disease can be divided into an acute and chronic phase based on its symptoms. In the chronic phase,
progressive heart failure is caused by the deterioration of heart muscle and the nervous system
(WHO, 2019; CDC, 2019). Within R. prolixus, T. cruzi exists in the highly infectious life stage
within the hindgut and is expelled during excretion after diuresis (Bern et al., 2011). Following
this excretion, T. cruzi can enter the host through bite wounds, or through mucous membranes
(CDC, 2019). Since there is no vaccine for Chagas disease, the most effective method of preventing
the spread of this disease is vector control (WHO, 2019). It is essential to examine the mechanisms
by which T. cruzi is spread by R. prolixus, which in turn will aid in the development of strategies
to decrease the spread of R. prolixus, and thereby the spread of Chagas disease.
Neuropeptides
Within nervous systems, there are several types of chemical messengers that function to
direct various essential processes that are critical for normal growth and development (Altstein &
Nässel, 2010). Amongst these chemical messengers are neuropeptides, that are synthesized and
released from neurons or neuroendocrine cells (Yeoh et al., 2017). They can function as
neuromodulators, neurotransmitters, or neurohormones (Orchard, 2009). This diversity in the
mode of action by which neuropeptides can function allows them to govern many physiological
processes. As neurotransmitters, their release is localized to the synaptic cleft, initiating rapid
changes via receptors on post-synaptic membranes. The effects of neurotransmitters are short term
3
as they are subject to degradation, diffusion, and reuptake from the synaptic cleft (Orchard, 2009).
As neurohormones, their signal is much more global as they are released from neurosecretory cells
into the circulatory system and act upon peripheral target tissues that express the receptor. These
effects are longer lasting since neurohormones typically function through G protein-coupled
receptors (GPCRs) resulting in downstream effects. Neuropeptides that do not fit within these
specified roles are termed neuromodulators, which includes any neuroactive compound that
modulates its target (Orchard, 2009; Schoofs et al., 2017). Bioactive neuropeptides are synthesized
from larger precursor molecules, known as prepropeptides. They are then targeted via the
regulatory secretory pathway to intracellular electron dense granules, where they are stored until
secretion (Elphick et al., 2018). Prepropeptides comprise of a signal peptide, propeptides (which
later become mature peptides), spacer peptides, and cleavage sites (Yeoh et al., 2017). The signal
peptide directs the prepropeptide to the secretory pathway, with monobasic and dibasic cleavage
sites surrounding the mature peptides, which are targets for various neuropeptidases (Yeoh et al.,
2017; Veenstra, 2000). Following processing, the neuropeptides can also be subject to post-
translational modifications, such as C-terminal amidation. (Yeoh et al., 2017). Mature
neuropeptides function by being released from granules and then typically binding onto specific
GPCRs on the cell membranes of target cells, initiating further downstream signaling pathways
(Elphick et al., 2018).
Within insects, and indeed other animals, neuropeptides represent the largest class of
chemical compounds involved in physiological processes such as development, reproduction,
metabolism, and behaviour (Altstein & Nässel, 2010). The first insect neuropeptide to be identified
was proctolin in Periplaneta americana, and there are currently approximately 50 identified
neuropeptide families in insects (Yeoh et al., 2017). Genes can encode for prepropeptides that vary
4
in the number of mature peptides, and in some cases the mature peptides may be structurally and
functionally distinct (Yeoh et al., 2017) An example of this is the Drosophila capability gene,
which encodes for two CAPA neuropeptides, and a third pyrokinin neuropeptide (Kean et al.,
2015).
G Protein-Coupled Receptors (GPCRs)
Neuropeptides function on their target tissues by binding onto their specified GPCRs,
resulting in the activation of a second messenger cascade (Grimmelikhuijzen & Hauser, 2012).
GPCRs, which comprise the largest group of membrane receptors, all share a similar structure
which includes an intracellular C-terminus, an extracellular N-terminus, and seven hydrophobic
transmembrane domains (TM1-TM7) (Gether, 2000; Pierce et al., 2002). These membrane
domains are linked by intracellular (ICL1-ICL3) and extracellular loops (ECL1-ECL3) (Gether,
2000). GPCRs can bind a variety of ligands, including peptides, biogenic amines, lipids, and
proteases (Gether, 2000). They can be classified into six classes (class A-F), with the possible
ligands varying within these classes. The rhodopsin-like (class A) family, which is the largest and
most studied family of GPCRs, includes receptors for various small molecules, neurotransmitters,
peptides and hormones (Munk et al., 2016). Some conserved regions of family A GPCRs include
the Asp-Arg-Tyr (DRY) motif, which is located on the intracellular side of TM3, and the Asp-Pro-
xx-Tyr (NPxxY) domain which is located within TM7 (Munk et al., 2016). The DRY (sometimes
DRH or ERY) motif is critical in inducing conformational changes that are required for receptor
activation, while the NPxxY domain is implicated in maintaining structural integrity (Fritze et al.,
2003; Capra et al., 2004; Rovati et al., 2007).
5
GPCRs are first synthesized within the endoplasmic reticulum (ER), and are then
transported to the cell membrane, which is its final target. During this transport, GPCRs undergo
various post-translational modifications to ensure biological activity (Duvernay et al., 2005).
GPCRs are often subject to phosphorylation within various sites at the C-terminus and intracellular
loops by protein kinases, which is essential for various mechanisms within GPCRs, such as
desensitization and internalization of the receptor (Ferguson, 2001; Tobin, 2008; Yang et al.,
2017). GPCRs are also targets of N-glycosylation, which is critical for trafficking and expression
to the cell surface and overall function (Michineau et al., 2005; Chen et al., 2010). Palmitoylation,
which is a lipid modification, involves the modification of Cys residues within the intracellular
loops, and C-terminus of GPCRs through the addition of a palmitic acid (Qanbar & Bouvier, 2003;
Goddard & Watts, 2002). This post-translational modification modulates various aspects of GPCR
function (Goddard & Watts, 2002).
GPCRs are activated through the binding of ligands, involving the function of
heterotrimeric G-proteins, which are a family of proteins made up of an α, β, and γ subunit.
Heterotrimeric G-proteins can be further divided into classes, depending on the type of α subunit.
Some examples are Gq, Gs, and Gi/o (Hamm, 1998; Caers et al., 2012). Gq induces the activation
of phospholipase C (PLC), activating the inositol triphosphate (IP3) pathway. This causes the
intracellular release of IP3 which binds onto an IP3 sensitive Ca2+ channel on the endoplasmic
reticulum and causing Ca2+ release. Ca2+ functions as a second messenger to further initiate various
downstream effects. Gs/Gi/o G-proteins are associated with adenyl cyclase (AC), an enzyme
responsible for the synthesis of cyclic adenosine monophosphate (cAMP) from adenosine
triphosphate (ATP) (Sadana & Dessauer, 2009; Caers et al., 2012). Gs is known to activate AC,
while Gi/o inhibits it (Hamm, 1998; Caers et al., 2012). Activation of the GPCR induces a
6
conformational change with its associated intracellular G-protein subunit, releasing guanosine
diphosphate (GDP), followed by the binding of guanosine triphosphate (GTP) to the α subunit of
the G-protein (Munk et al., 2016). The GTP-bound α subunit then dissociates from the receptor,
causing the release of the β-γ dimer. Both the GTP-bound α subunit and β-γ dimer can interact
with various intracellular mechanisms, eventually leading to the activation or inhibition of
signaling pathways. Investigating the structure and function of GPCRs provides value in the
development of next-generation pesticides, whereby neuropeptide GPCRs can be targeted in order
to disrupt critical processes within relevant species (Audsley & Down, 2015).
GPCRs have been observed to elicit their intracellular changes even in the absence of
ligand binding, known as constitutively active GPCRs. GPCR activity can be described by the
two-state model of GPCR activation, where GPCRs exist in two separate states, an inactive state
(R), and an active state (R*) (Seifert, 2002). The transition from R to R* state involves a
conformational change within the GPCR, which is induced by an agonist (Seifert, 2002). In
constitutively active GPCRs, transition to the R* state occurs independent of an agonist, stably
inducing basal G protein activity and its associated downstream effects (Steifert, 2002). Ligands
can function as full agonists that induce maximal activity of the constitutively active GPCR, or as
partial agonists that maintain the R* state of the GPCR to a lesser degree (Steifert, 2002). Inverse
agonists function to maintain the GPCR in its R state, in turn reducing basal G protein activity
(Steifert, 2002). Like agonists, inverse agonists can also function as full or partial inverse agonists
(Steifert, 2002). GPCR mutants causing constitutive activity or loss of constitutive activity have
been implicated in various diseases and are often used as pharmacological targets in drug
development (Chalmers & Behan, 2002).
7
Kinin
The kinin (or leucokinin) family of cephalomyotropic peptides was first identified through
high-performance liquid chromatography (HPLC) of head extracts of the cockroach, Leucophaea
maderae, and are known for their ability to stimulate hindgut contractions (Holman et al., 1986a,
b; Holman et al., 1987a, b). Following this, kinin-like neuropeptides were discovered in various
insect species, all sharing the C-terminal pentapeptide sequence FX1X2WG- amide where X1 can
be Ser, Phe, His, Asn, or Tyr and X2 can be Ser, Pro or Ala (Torfs et al., 1999). This core
pentapeptide sequence is essential for the biological activity of the neuropeptide (Nachman et al.,
1991; Coast et al., 1990). Kinins have been identified in various arthropods, with multiple isoforms
often being present within these species (Coast et al., 2002). Kinins also stimulate fluid secretion
from isolated Malpighian tubules (MTs) of Aedes aegypti females, suggesting a secondary diuretic
role of these neuropeptides (Hayes et al., 1989). In addition to their myotropic and diuretic effects,
kinins have also been implicated in a diverse set of functions, such as ecdysis-related behaviours
in Manduca sexta (Kim et al., 2006), and more recently, locomotor activity and metabolic rate in
Drosophila melanogaster (Zandawala et al., 2018).
Within R. prolixus, the Rhopr-kinin transcript encodes eighteen predicted kinins and
precursor associated peptides, the most found in any species (Table 1) (Te Brugge et al., 2011;
Bhatt et al., 2014). As seen in other species, Rhopr-kinins are primarily known for stimulating
hindgut contractions, with Rhopr-kinin 2 producing a potent effect (Bhatt et al., 2014). Kinins have
also been shown to have myotropic effects on the anterior midgut and salivary glands in R. prolixus
(Orchard & Te Brugge, 2002; Te Brugge et al., 2009). To observe the distribution of kinins within
R. prolixus, immunohistochemical analyses were performed, with kinin-like immunoreactive
staining observed in cell bodies and processes within the central nervous system (CNS), gut,
8
peripheral nerves, and peripheral neurons (Te Brugge et al., 2001). Kinins may also be implicated
in feeding-related behaviors within R. prolixus, due to their co-localization with the corticotropin-
releasing factor (CRF)-like diuretic hormone (Rhopr-CRF/DH). Rhopr-CRF/DH has been shown
to influence feeding as injection of Rhopr-CRF/DH into 5th instar insects resulted in the intake of
significantly smaller blood meals (Mollayeva et al., 2018). Kinin-like and CRF-like staining is
observed within the CNS, and endocrine cells of the midgut of R. prolixus, with a decrease in
staining observed in neurosecretory cells up to 2.5 hours after feeding. Levels of the kinin-like and
CRF-like staining are restored 1 day after feeding (Te Brugge et al., 1999; Te Brugge et al., 2001;
Te Brugge et al., 2002, Mollayeva et al., 2018).
The first kinin GPCR, belonging to the family A of GPCRs, was characterized in the snail,
Lymnaea stagnalis (Cox et al., 1997). Since then, kinin receptors have been functionally
characterized in a small number of insect species including D. melanogaster, A. aegypti, and
Anopheles stephensi (Radford et al., 2002; Radford et al., 2004; Pietrantonio et al., 2005). It is
hypothesized that all invertebrate kinin receptors may utilize intracellular Ca2+ as a second
messenger, as release of intracellular Ca2+ through activation of the IP3 pathway was observed in
L. Stagnalis, D. melanogaster, and A. aegypti (Cady and Hagedorn, 1999; Radford et al., 2002;
Pietrantonio et al., 2005). Currently, the kinin GPCR has not yet been characterized within R.
prolixus, therefore its downstream signaling pathway is unknown.
9
Table 1: Amino Acid sequences of Rhopr-kinins (K) processed from the kinin-precursor. Table
modified from (Te Brugge et al., 2011)
10
Neuropeptide name Neuropeptide sequence K-1 TNNRGNFAGNPRMRFSSWAa K-2 AKFSSWGa K-3 ANKFSSWAa
K-4 AKFSSWAa K-5 DEDRQKFSHWAa K-6 GAKFSSWAa
K-7 AKFNSWGa K-8 LSINPWKKIDDNGa K-9 AKFSSWGa K-10 ADDDWLKKARFNSWGa
K-109–15 ARFNSWGa K-11 SAAAYTPLSWKRKPIFSSWGa K-111–11 SAAAYTPLSW-OH
K-1113–20 KPIFSSWGa K-1114–20 PIFSSWGa K-12 RGPDFYAWGa
K-122–9 GPDFYAWGa KPP-6 FSNEFMNDNNDIEKNIVEE-OH
11
CAPA
CAPA neuropeptides, a family of neuropeptides belonging to the PRXamide superfamily
are characterized by their consensus carboxyl terminal sequence A/PFPRV-NH2 (Paluzzi, 2012;
Alford et al., 2019a). The first CAPA neuropeptide was originally isolated in ventral nerve chord
extracts of M. sexta (Huessman et al., 1995). Initially named CAP2b, this neuropeptide was found
to increase heart rate in M. sexta, hence it’s designation as a cardioactive peptide (CAP). CAP2b is
one of the many cardioactive peptides isolated from M. sexta (CAP1a, CAP1b, CAP2a, CAP2b), but
shares no sequence homology, making it a novel neuropeptide (Tublitz & Truman, 1985a–d;
Huessman et al., 1995). CAP2b was also found to have diuretic effects, as it increased fluid
secretion within D. melanogaster (Huessman et al., 1995). Separately from CAP2b, a myotropic
neuropeptide was isolated from the perisympathetic organs of P. americana, named
periviscerokinin (Pea-PVK) (Predel et al., 1995). Due to structural similarities between CAP2b and
periviscerokinin, they can be grouped together into the CAPA family of neuropeptides (Predel &
Wegener, 2006). Following sequencing of the CAPA gene in M. sexta, CAP2b was later named
ManseCAPA-1 (Loi & Tublitz, 2004). In D. melanogaster, a gene named capability was identified
that encoded two ManseCAPA-1 related peptides (capa-1, capa-2), and a third pyrokinin-related
neuropeptide (Kean et al., 2015). Like ManseCAPA-1, capa-1 and capa-2 also stimulated MT fluid
secretion within D. melanogaster (Kean et al., 2015).
As seen in other insects, the CAPA neuropeptides within R. prolixus are encoded by the
RhoprCAPA transcript, named RhoprCAPA-1 (SPISSVGLFPFLRA-NH2), RhoprCAPA-2
(EGGFISPRV-NH2), and Rhopr-pk1 (NGGGGNGGGLWFGPRL-NH2) (Paluzzi et al., 2008).
RhoprCAPA-1 lacks the characteristic PRVamide sequence found in other CAPA neuropeptides
(Paluzzi et al., 2008). A second CAPA gene paralog was identified in R. prolixus that shares a very
12
high sequence similarity with the first RhoprCAPA transcript, named RhoprCAPA-β (Paluzzi &
Orchard, 2010). The CAPA neuropeptides encoded by RhoprCAPA-β are named RhoprCAPA-
β1, RhoprCAPA-β2, and RhoprCAPA-βpk1. As a second gene paralog for the RhoprCAPA
transcript was identified, the first transcript is referred to as RhoprCAPA-α, with the encoded
neuropeptides renamed to the following: RhoprCAPA-α1, RhoprCAPA-α2, and Rhopr-αpk1
(Paluzzi et al., 2009). CAPA neuropeptides are distributed in various neurons within the nervous
system of R. prolixus, including the brain, suboesophageal ganglion (SOG), prothoracic ganglion,
and in the abdominal neuromeres of the mesothoracic ganglionic mass (MTGM) (Paluzzi et al,
2006; Paluzzi & Orchard, 2010; Paluzzi et al., 2008). RhoprCAPA-α2 was identified as the first
anti-diuretic hormone within R. prolixus, as it inhibited serotonin [5-hydroxytryptamine (5-HT)]
stimulated effects on the MT, and anterior midgut (Paluzzi et al., 2006; Ianowski et al., 2009). The
effects of RhoprCAPA-α2 may be related to post-feeding diuresis, as immunohistochemical
staining of three pairs of neurosecretory cells within the abdominal neuromeres is significantly
decreased three to four hours post-blood meal (Paluzzi et al., 2006).
CAPA receptors, belonging to family A GPCRS, have been characterized in insects such
as D. melanogaster, A. gambiae, and Tribolium castaneum (Iversen et al., 2002; Olsen et al., 2007;
Jiang et al., 2015). The R. prolixus CAPA receptor (capa-r) has been characterized, and it was
identified as the first antidiuretic hormone receptor in insects, with cyclic guanosine
monophosphate (cGMP) likely acting as a second messenger (Paluzzi et al., 2010). Expression of
capa-r is highest in the MTs and anterior midgut of R. prolixus, which are known targets of
RhoprCAPA-α2 (Paluzzi et al., 2010). Expression of capa-r has also been confirmed on the
hindgut, suggesting a role of CAPA neuropeptides on this tissue (Paluzi et al., 2010). Two
transcript variants of capa-r were identified, named capa-r1 and capa-r2, however capa-r2
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encodes an atypical GPCR (Paluzzi et al., 2010). As RhoprCAPA- α1 lacks the characteristic
PRVamide sequence, it is unable to activate the capa-r1 receptor (Paluzzi et al., 2010).
Post-Feeding Physiology
Feeding within R. prolixus has been a widely studied phenomenon, due to the dependence
of a blood meal to successfully transition to the next instar and to initiate many crucial
developmental and reproductive processes. The feeding strategy of R. prolixus can add a great deal
of osmotic stress, as feeding on such a large blood meal results in the consumption of excess water
and salts. This large blood meal also causes a large increase in the body weight of the insect,
leaving it in a state of susceptibility to predation (Orchard, 2006; Orchard, 2009). The blood meal
serves as a signal to initiate short-term endocrinological and physiological changes to lower the
insect’s mass and return to a homeostatic state (Maddrell, 1976).
Diuresis
Within the digestive tract of R. prolixus, the anterior midgut, upper and lower MTs are the
tissues primarily responsible for the rapid absorption and secretion of water and salts, resulting in
the production of primary urine (Coast et al., 2002; Maddrell, 1969). Once blood has entered the
midgut, absorption of water and NaCl occurs from the anterior midgut into the haemolymph.
Following this, the upper MTs secrete a fluid containing a high NaCl and KCl content. The final
modification of this fluid occurs by the lower MTs, which reabsorb KCl, resulting in the production
of primary urine (Orchard, 2009). This ionic movement that occurs within the anterior midgut and
MTs is fast-acting, occurring within minutes following feeding (Maddrell, 1976; Orchard, 2009).
The cells in the epithelium of MTs within R. prolixus have been described as “the fastest fluid
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secreting cells known”, further emphasizing how quickly post-feeding diuresis occurs to rid the
insect of excess fluids (Maddrell, 1991).
The absorption and secretion mediated by the anterior midgut and MTs occurs in a highly
timed and coordinated manner, and these tissues are under direct control by a host of diuretic and
antidiuretic factors (Coast et al., 2002). The diuretic neurohormones 5-HT, CRF/DH, and
calcitonin-like diuretic hormone (Rhopr-DH31) are primarily responsible for the diuretic action of
the anterior midgut and the MTs (Maddrell et al., 1971; Te Brugge et al., 2011, Te Brugge et al.,
2005), with evidence of their release observed through immunohistochemical staining and analysis
of haemolymph composition. There is a reduction in 5-HT-like immunoreactive staining in central
and peripheral tissues during feeding in 5th instar insects, with a rapid rise in serotonin within the
haemolymph (Lange et al., 1988; Orchard et al., 1989). A decrease of CRF-like and kinin-like
immunoreactive staining in neurosecretory cells is seen up to 2.5 hours after feeding, with levels
being restored 1 day after feeding (Te Brugge et al., 1999; Te Brugge et al., 2001; Te Brugge et
al., 2002; Mollayeva et al., 2018). In addition, there is a reduction in DH31-like staining within
nerve processes post-feeding (Te Brugge et al., 2005). These reductions in immunoreactive
staining post-feeding is likely due to the release of these neurohormones during post-feeding
diuresis.
5-HT increases the rate of fluid transport across the anterior midgut and increases the
frequency of contractions in both in vitro and in vivo studies within 5th instar R. prolixus (Te
Brugge et al., 2009; Barrett et al., 1993). Within the MTs, 5-HT has been shown to stimulate upper
MT fluid secretion and stimulates reabsorption within the lower MTs (Te Brugge et al., 2009;
Donini et al., 2008). In the upper and lower MTs, 5-HT also increases cAMP content, which
15
suggests that cAMP is acting as a second messenger to facilitate this process (Te Brugge et al.,
2002). When tested on the anterior midgut, Rhopr-DH31 failed to have any effects on fluid
absorption but was found to increase the frequency of contractions and cAMP levels (Te Brugge
et al., 2009). Within the MTs, Rhopr-DH31 only induces minor increases in fluid secretion in the
upper MTs but has no effect on the lower MTs (Te Brugge et al., 2002; Donini et al., 2008). Rhopr-
CRF/DH stimulates fluid transport and has myotropic effects within the anterior midgut, and
stimulates upper MT fluid secretion (Te Brugge et al., 2002; Te Brugge et al., 2009). However,
within the lower MTs, Rhopr-CRF/DH does not have any effects on reabsorption (Donini et al.,
2008). An increase in cAMP levels is also exhibited within these tissues following treatment with
Rhopr-CRF/DH (Te Brugge et al., 2002). Within the MTs, it is suggested that there is a synergistic
interaction between Rhopr-CRF/DH and 5-HT to increase fluid secretion rates, highlighting the
coordination required during post-feeding diuresis (Maddrell et al., 1971; Barrett & Orchard,
1990). Following the production of primary urine, the termination of diuresis occurs to prevent
any further fluid loss (Maddrell, 1964). As found within diuresis, this termination is under the
control of the anti-diuretic hormone, RhoprCAPA-2 (Paluzzi et al., 2008; Orchard & Paluzzi,
2009). RhoprCAPA-2 inhibits 5-HT stimulated fluid secretion in the upper MTs, and inhibits 5-
HT-stimulated fluid absorption in the anterior midgut (Paluzzi, 2006; Paluzzi et al., 2008; Ianowski
et al., 2009)
Excretion
Following the production of primary urine, the final step occurs within the hindgut,
whereby contractions of the hindgut result in excretion of this urine (Maddrell, 1964). In addition
to its role in post-feeding diuresis, the hindgut is a tissue of epidemiological relevance, as the T.
cruzi parasite exists in its highly infective stage in the hindgut, and so hindgut contractions result
16
in the transmission of the parasite along with urine onto the host (Bern et al, 2011). The myotropic
action of the hindgut is under control by many neuroendocrine factors such as 5-HT, Rhopr-DH31,
Rhopr-CRF/DH, kinins and tachykinins (Te Brugge et al., 2002; Bhatt et al., 2014; Haddad et al.,
2018). As seen in the MTs, interactions between different neuropeptide families within the hindgut
has been observed, with kinins, tachykinins and CRF/DH exhibiting additive or co-operative
effects on hindgut contraction stimulation (Bhatt et al., 2014; Haddad et al., 2018).
Neuropeptide Analogs
From an agrochemical and medical perspective, neuropeptide signaling is a field of interest,
as many critical physiological processes and behaviors are under direct control of neuropeptides.
They can be used in insecticidal strategies to interfere with the normal functioning of pests and
disease vectors, thereby preventing the detrimental effects of these insect species (Gäde &
Goldsworthy, 2003). As many insects have developed resistance to many traditional insecticides,
neuropeptides provide a promising alternative to combat particularly damaging species (Nachman
& Smagghe, 2011). Neuropeptides have been studied as lead compounds in the development of
more environmentally friendly pest control strategies, due to their specificity and activity at low
concentrations (Nachman & Smagghe, 2011). Some of the limitations with the use of native
neuropeptides is the susceptibility to a variety neuropeptidases within the insect’s haemolymph
and tissues, which results in the inactivation of the neuropeptide, reducing its bioavailability. In
addition, neuropeptides in their native form are unable to penetrate an insect’s exoskeleton, thus
severely limiting the possible forms of delivery (Isaac et al., 2009; Menn & Bořkovec, 1989). The
study of neuropeptides, their receptors and the biochemical features required for successful
interaction has allowed the development of compounds that function as agonists or antagonists to
specific neuropeptide receptors (Menn & Bořkovec, 1989; Keeley & Hayes, 1987). These
17
compounds, known as neuropeptide analogs, are synthesized with modifications to their amino
acid sequence to overcome the limitations associated with native neuropeptides, so they can be
successfully used in pest control strategies (Nachman, 2009). Various analogs of many well-
studied neuropeptide families have been synthesized (eg. kinins, CAPA) and have been examined
for their effects on tissues within several insect species (Bhatt et al., 2014; Lange et al., 2016;
Smagghe et al., 2010; Alford et al., 2019a; Alford et al., 2019b)
Within kinins, the C-terminal pentapeptide core region between the Ser (or Pro) and
conserved Trp residues is susceptible to primary hydrolysis. A secondary site of hydrolysis is also
found outside of the core region, at the neuropeptide bond N-terminal to Phe (Nachman et al.,
1997a; Nachman et al., 1997b). The fly angiotensin converting enzyme (ACE) can cleave the
primary hydrolysis site, with neprilysin (NEP) cleaving the primary and secondary hydrolysis sites.
Replacement of the Ser (or Pro) with an Aib residue blocks ACE or NEP hydrolysis, while
mimicking a critical conformation required for activity (Nachman et al., 1997a; Nachman et al.,
1997b; Nachman et al., 2002; Xiong et al., 2018). The Aib-kinin analog induces potent changes in
physiology, and in some cases disrupt essential processes. Within the aphids Myzus persicae and
Macrosiphum rosae, decreased survival was exhibited under cold stress exposure after kinin
analog treatment. In Acyrthosiphon pisum, the kinin analog induced antifeedant activity and high
mortality (Smagghe et al., 2010). Aib-containing kinin analogs also induced physiological changes
within R. prolixus, as they were found to have antifeedant effects, and disrupted ecdysis (Lange et
al., 2016). Aib-containing analogs had potent myotropic effects in R. prolixus, as they induced
stronger hindgut contractions than native Rhopr-kinins (Bhatt et al., 2014).
18
More recently, analogs from the PRXamide family of neuropeptides have been
synthesized, with addition of hydrophobic moieties to the N-terminus to increase greater in vivo
stability (Jurenka, 2015; Zhang et al., 2011). Second generation analogs for CAPA neuropeptides
have also been synthesized with steric hinderances adjacent to the C-terminal position, thus biasing
it’s binding to CAPA receptors, since cross-reactivity has been exhibited on CAPA receptors with
CAPA and pyrokinin neuropeptides (Jiang et al., 2015; Paluzzi et al, 2010; Paluzzi & O’Donnell,
2012). Within D. melanogaster and D. suzukii, flies microinjected with a CAPA analog had an
increased survival rate under desiccation stress (Alford et al., 2019a). In the aphids M. persicae
and M. rosae, the CAPA analogs accelerated aphid mortality under desiccation and starvation
stress, with the analog causing enhanced mortality under cold stress in M. persicae (Alford et al.,
2019b). The ability of these neuropeptide analogs to disrupt physiological processes, and in some
cases cause mortality within insects highlights their promise in pest control strategies. It is
imperative to continue investigating the use of neuropeptide analogs to prevent the further spread
of pests and disease vectors.
Objectives of Thesis
Within R. prolixus, feeding and diuresis are epidemiologically-relevant behaviours, as they
are implicated in the transmission of the T. cruzi parasite. Previous studies have investigated the
role of kinins and CAPA, with Rhopr-kinins primarily exerting myotropic effects on target tissues,
and RhoprCAPA-2 functioning as an anti-diuretic hormone. Kinin and CAPA neuropeptides will
be further investigated on feeding and diuresis-related tissues, which will provide insight into their
roles as neuroendocrine factors in R. prolixus. Neuropeptide analogs have been developed for
Rhopr-kinin and RhoprCAPA-2, that may influence processes related to disease transmission.
These analogs will be investigated for their efficacy in inducing changes in physiology, to
19
determine whether they are lead compounds for the development of pesticides. The physiological
approaches to investigate these neuropeptides and analogs are listed below:
• Analyze the in vivo effects of injected kinin, CAPA, and their analogs on feeding and
post-feeding diuresis.
• Determine the effects of Rhopr-kinin 2, the Aib-containing kinin analog 2139[Ф1]wp-
2, CAPA neuropeptides, and CAPA analog 2129-SP3[Ф3]wp-2 on the hindgut of R.
prolixus.
• Determine the effects of 2129-SP3[Ф3]wp-2 on fluid secretion in the MTs of R.
prolixus.
Currently, the R. prolixus kinin receptor has not yet been characterized. Identifying,
cloning, and sequencing the Rhopr-kinin receptor will aid in the development of next-generation
pest control strategies. The molecular approach to characterizing this receptor is listed below:
• Isolate, clone and sequence the Rhopr-kinin GPCR.
• Analyze and predict the biochemical features and structural characteristics of the
kinin GPCR using online tools.
• Determine the spatial expression of the kinin receptor transcript within the CNS and
tissues involved in feeding, diuresis and excretion.
• Target the kinin receptor through RNA interference (RNAi) and examine effects of
knockdown through hindgut contraction assays and feeding bioassays.
20
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Chapter 2 Physiological Effects of Biostable kinin and CAPA Analogs in the
Chagas Disease Vector, Rhodnius prolixus
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Abstract
In the Chagas disease vector Rhodnius prolixus, the kinin and CAPA family of
neuropeptides are implicated in feeding and diuresis-related behaviours, with Rhopr-kinins
stimulating contractions of the midgut, salivary glands, and hindgut, with RhoprCAPA-2
functioning as an anti-diuretic hormone. The current study examined the effects of kinin and
CAPA neuropeptides and their analogs on feeding and diuresis, and on hindgut contractions and
MT fluid secretion in R. prolixus. The biostable Aib-containing kinin analog 2139[F1]wp-2 was
found to have antifeedant effects, and to be more potent than Rhopr-kinin 2 in stimulating hindgut
contractions. The CAPA analog 2129-SP3[F3]wp-2 induced the intake of a larger blood meal, and
increased the rate of post-prandial rapid diuresis. RhoprCAPA-2, but not its analog, potentiated
hindgut contractions induced by Rhopr-kinin 2. Potentiation was observed with the CAPA analog
on 5-HT-stimulated increases in frequency of hindgut contractions, whereas RhoprCAPA-2
inhibited this 5-HT-mediated stimulation. The CAPA analog induced hindgut contractions and
prevented the inhibition induced by RhoprCAPA-2 on 5-HT-stimulated MT secretion. These
results demonstrate novel interactions between Rhopr-kinin and RhoprCAPA-2 on the hindgut,
possibly influencing post-feeding excretion. The kinin analog is a potent agonist of the kinin
receptor, and the CAPA analog an antagonist of the CAPA receptor. The use of neuropeptide
mimetics is a promising approach to vector control as they can disrupt behaviours, and the effects
of these neuropeptide analogs highlight their value as lead compounds, given their ability to
interfere with epidemiologically-relevant behaviours.
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Introduction
The blood-gorging hemipteran Rhodnius prolixus is a domestic vector of Chagas disease
in Central and South America, caused by the transmission of Trypanosoma cruzi, a flagellate
parasite (Moncayo, 2003; Dujardin et al., 1998). R. prolixus consumes a blood meal that can be up
to 10 times its body weight during each instar, following which the insect rids itself of the excess
water and salt from the blood meal. The transmission of the parasite to its host occurs during this
post-prandial diuresis, and so an understanding of the physiology of feeding and diuresis-related
behaviours is essential in understanding disease transmission (Orchard, 2006; Orchard & Paluzzi,
2009). The systems controlling diuresis include the Malpighian tubules (MTs), midgut and
hindgut, which are subject to control by various diuretic and antidiuretic hormones and myotropic
factors (Coast et al., 2002; Orchard, 2006; Orchard & Paluzzi, 2009).
Many essential processes within insects are regulated by neuropeptides which act upon
target tissues through their release as neurohormones into the haemolymph or as neuromodulators
(Schoofs et al., 2017). Insect kinins, first isolated from Leucophaea maderae head extracts, share
the C-terminal pentapeptide sequence FX1X2WG-amide (where X1 can be Ser, Phe, His, Asn, or
Tyr and X2 can be Ser, Pro or Ala) (Holman et al., 1986a, b; Holman et al., 1987a, b). Kinins have
been implicated in various behaviours, such as hindgut contraction (Bhatt et al., 2014), MT fluid
secretion (O’Donnell et al., 1998; Terhzaz et al., 1999; Rosay et al., 1997), ecdysis (Kim et al.,
2006), and more recently locomotor activity and metabolic rate (Zandawala et al., 2018). The R.
prolixus kinin (Rhopr-kinin) transcript encodes for eighteen predicted kinins and precursor
associated peptides (Te Brugge et al., 2011a; Bhatt et al., 2014) and these kinins stimulate R.
prolixus hindgut, anterior midgut, and salivary gland contractions (Orchard & Te Brugge, 2002;
Te Brugge et al., 2009; Te Brugge et al., 2011; Bhatt et al., 2014). Interestingly, co-localization of
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Rhopr-kinin has been shown with corticotropin-releasing factor (CRF)-like diuretic hormone
(Rhopr-CRF/DH) within the R. prolixus central nervous system (CNS) and endocrine cells of the
midgut. A decrease of CRF-like and kinin-like immunoreactive staining in neurosecretory cells is
seen up to 2.5 hours after feeding, with levels being restored 1 day after feeding, suggesting a role
of CRF and kinin in feeding-related behaviours (Te Brugge et al., 1999; Te Brugge et al., 2001;
Te Brugge et al., 2002; Mollayeva et al., 2018). In the R. prolixus excretory system, the MTs play
a dominant role in the regulation of urine volume and composition (Coast, 2009), primarily under
control by the diuretic neurohormones serotonin [5-hydroxytryptamine (5-HT)], calcitonin-like
diuretic hormone (DH31), and Rhopr-CRF/DH (Maddrell, 1971; Te Brugge et al., 2005; Te Brugge
et al., 2011b), and the antidiuretic hormone R. prolixus CAPA (Rhopr-CAPA) (Orchard & Paluzzi,
2009; Ianowski et al., 2009; Paluzzi et al., 2008). CAPA peptides are encoded by the capability
genes, initially identified in Drosophila melanogaster (Kean et al., 2002). The first two CAPA
peptides encoded by the gene contain the consensus carboxyl terminal sequence A/PFPRV-NH2,
with the third peptide containing the consensus carboxyl terminal sequence G/MWFGPRL-NH2,
typically referred to as a pyrokinin (PK)-related peptide (Paluzzi, 2012; Paluzzi et al., 2008).
CAPA peptides were originally discovered in Manduca sexta for their cardioacceletatory effects
and display diuretic, anti-diuretic, and myotropic effects in a variety of species (Huesmann et al.,
1995; Predel & Wegener, 2006; Paluzzi et al., 2008). The RhoprCAPA transcript encodes three
CAPA peptides, as found in other species: RhoprCAPA-1, RhoprCAPA-2, and RhoprCAPA-pk1.
RhoprCAPA-2 has been found to inhibit 5-HT-stimulated secretion by the MTs and absorption
from anterior midgut (Ianowski et al., 2010; Orchard & Paluzzi, 2009; Paluzzi et al., 2008). The
effects of the CAPA peptides are mediated by the RhoprCAPA G protein-coupled receptor
(GPCR) (capa-r) (Paluzzi et al., 2010). Expression of the capa-r transcript has been confirmed in
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the hindgut of R. prolixus, suggesting that CAPA peptides may influence hindgut contractions
(Paluzzi et al., 2010).
From an agrochemical and medical perspective, neuropeptides and neuropeptide analogs
are compounds of interest for the disruption of critical functions as a means of pest control (Jiang
et al., 2015). Neuropeptide analogs have been synthesized with a modified chemical structure in
order to overcome limitations associated with delivery of the compound (eg. movement through
the cuticle, endopeptidases, exopeptidases) (Nachman et al., 1997). In Musca domestica, addition
of alpha-aminoisobutyric acid (Aib) to a kinin analog resulted in resistance to hydrolysis by
angiotensin converting enzyme (ACE) and neprilysin (NEP) and the analog was found to be potent
in inducing myotropic activity in L. maderae hindguts (Nachman et al., 1997). Recently, the kinin
analog 2139 was shown to stimulate fluid secretion in D. melanogaster, and significantly reduced
survival under desiccation stress (Alford et al., 2019a). Decreased survival was also exhibited in
Myzus persicae and Macrosiphum rosae under cold stress exposure after kinin analog treatment
(Alford et al., 2019b). In R. prolixus, an Aib-containing kinin analog was found to have antifeedant
effects, as insects only consumed 60% of a blood meal that contained the analog (Lange et al.,
2016).
The CAPA analog 2129-SP3[F3]wp-2 has been designed with the addition of hydrophobic
moieties to the N-terminus to increase greater in vivo stability, and also possesses a steric
hinderance adjacent to the alpha carbon in the C-terminal position, directing its binding to a CAPA
receptor whilst interfering with any PK receptor binding (Zhang et al., 2011; Jurenka, 2015; Alford
et al., 2019a). This CAPA analog also influences desiccation and starvation survival, as it
significantly improved D. suzukii desiccation survival, while significantly increasing the
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desiccation and starvation mortality in M. persicae and M. rosae (Alford et al., 2019a; Alford et
al., 2019b).
In this study, we examine the effects of the biostable Aib-containing kinin analog
2139[F1]wp-2 and an antagonist of the CAPA receptor, CAPA analog 2129-SP3[F3]wp-2 (Table
1) on feeding and diuresis-related behaviours, including changes in blood meal size, hindgut
contractions, and excretion rate. Due to the presence of CAPA receptors on the hindgut (Paluzzi
et al., 2010), we also further investigated the role of members of the RhoprCAPA family of
peptides on hindgut contractions. Examining the effects of the kinin and CAPA analogs on the
physiology of R. prolixus will assist in determining the potential value of these analogs in wide
scale pest control strategies.
Materials and Methods
Animals
5th instar male and female R. prolixus were obtained from an established colony at the
University of Toronto Mississauga. Insects were reared at 25oC and 50% humidity in incubators,
and were fed defibrinated rabbit blood (Cedarlane Laboratories, Burlington, ON, Canada) once in
each instar. Tissues were dissected from 5th instar R. prolixus 3-5 weeks post-feeding as 4th instars.
All tissue dissections were performed in R. prolixus physiological saline, consisting of 150 mM
NaCl, 8.6 mM KCl, 2.0 mM CaCl2, 4.0 mM NaHCO3, 8.5 MgCl2 , 0.02 mM HEPES and 34 mM
glucose in pH 7.0.
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Chemicals
Rhopr-kinin 2, RhoprCAPA-1, RhoprCAPA-2, and RhoprCAPA-pk1 were custom
synthesized by Genscript (Piscataway, NJ, USA). The peptides were then reconstituted in double-
distilled water into stock solutions at 10-3 M and stored at -20oC. Stock solutions of the Aib-
containing insect kinin analog (2139[F1]wp-2) with the amino acid sequence Phe-Phe-Aib-Trp-
Gly-NH2 (Nachman et. al., 1997), and insect CAPA analog (2129-SP3[F3]wp-2) with the
sequence 2Abf-Suc-ATPRIa synthesized as previously described (Nachman et al., 1997; Alford et
al., 2019b) were prepared in 80 % aqueous acetonitrile (ACN) containing 0.01% trifluoroacetic
acid (TFA), and stored at 4oC at a concentration of 10-3 M. Peptides and analogs were diluted in
physiological saline to various concentrations to be used during physiological assays
Hindgut Contraction Assay via Force Transducer
The R. prolixus hindgut was isolated under physiological saline, along with the cuticle at
the posterior end and fixed onto a Sylgard-coated dish using minuten pins through the cuticle and
bathed in 200 μl of physiological saline. One end of a fine silk thread was tied to the anterior end
of the hindgut, with the other end tied to a Grass FT03 force transducer (Astro-Nova Inc., Rhode
Island, USA). The amplitude of basal tonus changes were recorded using the PicoScope 2204
Oscilloscope (Pico Technology, Cambridgeshire, UK). Tissues were equilibrated in saline for 10
minutes. Peptides, including their analogs, were applied by addition of 100 μl of various
concentrations of the peptides in saline concurrent with removal of 100 μl of the bath saline to
ensure the bath volume remained constant. The preparations were washed with saline between test
doses of peptide and the bath volume was maintained at 200 μl of saline. The recorded traces were
analyzed for changes in basal tonus.
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Hindgut Contraction Assay via Impedance
The R. prolixus hindgut was isolated under physiological saline, along with the cuticle at
the posterior end and fixed onto a Sylgard-coated dish using minuten pins through the cuticle and
the anterior end and bathed in 200 μl of physiological saline. Electrodes were placed on either side
of the anterior region of the hindgut. Peristaltic contractions were monitored through a UFI
impedance converter (Model 2991, Morro Bay, CA, USA). The frequency of the hindgut
contractions was recorded using the PicoScope 2204 Oscilloscope (Pico Technology,
Cambridgeshire, UK). Tissues were equilibrated with saline for 10 minutes. Peptides and analogs
were applied onto the tissue by addition of 100 μl of various concentrations in saline concurrent
with removal of 100 μl of the saline, ensuring that the volume of saline within the bath remained
constant. To validate the recorded contractions, the tissue was observed visually to correlate which
deflections represent contractions. The frequency of contractions was measured for 3 minutes, and
the recorded traces were analyzed.
Feeding Bioassay
Unfed 5th instar insects (3-5 weeks post feeding as 4th instars) were separated into 3 groups
of 20 with similar average weights. Each group was injected through the membrane at the junction
of the hind leg with the abdomen with one of the following: 1 μl Rhopr-kinin 2 (10-4 M), 1 μl kinin
analog 2139[F1]wp-2 (10-4 M), 1 μl RhoprCAPA-2 (10-4 M), 1 μl CAPA analog 2129-
SP3[F3]wp-2 (10-4 M), or 1 μl physiological saline. After a 2-hour recovery period, each group
was placed in a 10 cm diameter glass jar and fed on 20 mL of warm defibrinated rabbit blood for
20 minutes. Insects from each group were individually weighed immediately after feeding (time
0) and maintained in individual cubicles. Weights were later recorded at 1, 2, 3, and 4-hour time
points. Since 5th instar R. prolixus tend to take a blood meal 8-10 times their initial body weight,
37
insects that fed less than 1 times their body weight were considered not fed and excluded from the
data, as were those punctured during the weighing process.
Malpighian Tubule Secretion Assay
Whole MTs from 5th instar insects were dissected under saline using glass probes and
transferred to a Sylgard-coated Petri dish containing 20 µl drops of saline overlaid with water-
saturated mineral oil. The proximal end of the tubule was pulled out of the saline and wrapped
around a minuten pin. Excess tubule from the proximal end was cut prior to wrapping, and the
tubules were nicked gently at the pin. The equilibrating saline was removed and replaced with 10-
8 M 5-HT (Sigma, Oakville, ON, Canada), the CAPA analog 2129-SP3[F3]wp-2, a mixture of 10-
8 M 5-HT and 10-7 M RhoprCAPA-2, or a mixture of 10-8 M 5-HT, 10-7 M RhoprCAPA-2 and
different concentrations of the CAPA analog 2129-SP3[F3]wp-2. Tubules were allowed to secrete
for 30 minutes. Droplets of secreted fluid from the nicked end of the tubule were then collected
using an oil filled micropipette tip, and the diameter of the droplet was measured using an eyepiece
micrometer on the bottom of the Sylgard-coated Petri dish. The droplet volume was then calculated
using the equation V=(π/6)d3 where d is the diameter of the droplet measured. At the end of the
experiments, tubules were stimulated with 10-6 M 5-HT and the maximal rate of secretion was
measured to check viability of the tissues.
Results
In Vivo Effects of Kinin and CAPA Analogs on Feeding and Diuresis
Injection of Rhopr-kinin 2 prior to feeding did not alter the size of blood meal consumed
over a 20-minute feed as compared to saline injected insects (Fig. 1A). On the other hand, injection
of the kinin analog 2139[F1]wp-2 prior to feeding led to a significant decrease in the size of the
38
blood meal consumed over a 20 minute period (Fig. 1A). Rapid post-feeding diuresis occurs over
the subsequent 3-4 hours and this can be monitored by measuring the loss of weight due to
excretion over time. Rhopr-kinin 2 and the kinin analog 2139[F1]wp-2 did not alter the rate of
diuresis over 4 hours (Fig. 1B,1C).
The effects of RhoprCAPA-2 and the CAPA analog, 2129-SP3[F3]wp-2, were also
examined on feeding and rate of diuresis in 5th instar insects. Injection of RhoprCAPA-2 had no
effect on the size of the blood meal, whereas injection of the CAPA analog 2129-SP3[F3]wp-2,
resulted in a larger blood meal being consumed (Fig. 2A). Injection of RhoprCAPA-2 did not
influence the rate of diuresis over 4 hours (Fig. 2B); however, the CAPA analog, 2129-
SP3[F3]wp-2, which resulted in a larger blood meal taken appeared to have an increased rate of
diuresis over the first hour after feeding (Fig. 2C).
In Vitro Effects of Kinin and CAPA Analogs on Hindgut Contractions
To further investigate the analogs, we turned to in vitro preparations of tissues associated
with feeding, namely the hindgut and MTs. Both Rhopr-kinin 2 and the kinin analog, 2139[F1]wp-
2, resulted in dose-dependent increases in basal tonus of the hindgut with threshold at 10-10 M for
Rhopr-kinin 2 and 10-14 M for 2139[F1]wp-2, illustrating the potency of the analog (Fig. 3). In
addition, 2139[F1]wp-2 induced stronger contractions than Rhopr-kinin 2. The EC50 value of
2139[F1]wp-2 is approximately 5.5 x 10-10 M, whereas the EC50 value for Rhopr-kinin 2 is
approximately 5.5 x 10-9 M. (Fig. 3C, 3D) The effects of 2139[F1]wp-2 were more difficult to
reverse and required more washes in saline than Rhopr-kinin 2. Neither RhoprCAPA-1 (not
shown), RhoprCAPA-2 (Fig. 4A) nor RhoprPK-1 (not shown) altered contractions of hindgut.
39
Interestingly, the CAPA analog, 2129-SP3[F3]wp-2, stimulated dose-dependent increases in
hindgut contractions (Fig. 4B, 4C). The threshold is at 10-10 M, EC50 of approximately 10-8 M, and
maximum tension at 10-6 M (Fig. 4C). In order to examine for any interaction between Rhopr-
kinin 2 and RhoprCAPA-2, the two peptides were applied simultaneously on the hindgut (Fig. 5,
6). Application of Rhopr-kinin 2 along with RhoprCAPA-2 resulted in statistically significant
increases in hindgut contractions relative to Rhopr-kinin 2 alone (Fig. 5A-5C). Interestingly, this
potentiation of hindgut contractions was not observed with the CAPA analog, 2129-SP3[F3]wp-
2 (Fig. 6).
To further examine any co-operative effects of RhoprCAPA-2 and the CAPA analog 2129-
SP3[F3]wp-2, changes in 5-HT-stimulated increases in the frequency of hindgut contractions were
measured. As the contractions induced by 5-HT are not easily monitored by the force transducer
(Te Brugge et al., 2002; Bhatt et al., 2014), an impedance monitor was used to assess changes in
the frequency of hindgut contractions. Varying concentrations of RhoprCAPA-2 and 2129-
SP3[F3]wp-2 were each applied simultaneously with 10-8 M 5-HT on the hindgut (Fig. 7, 8). 2129-
SP3[F3]wp-2 potentiated the effects of 10-8 M 5-HT, with a statistically significant increase in
hindgut frequency observed at a concentration of 10-7 M 2129-SP3[F3]wp-2 (Fig. 7B, 7C).
However, Rhopr-CAPA 2 was found to inhibit this 5-HT-mediated increase in frequency, with a
statistically significant reduction in frequency at 10-6 M (Fig. 8B, 8C).
In Vitro Effects of CAPA Analogs on Malpighian Tubule Secretion
To determine the effects of the CAPA analog 2129-SP3[F3]wp-2 on diuresis in vitro,
varying concentrations of the analog were tested on unstimulated tubules, and 5-HT-stimulated
tubules. The analog had no effect on unstimulated tubules (not shown) and failed to have any
40
potentiation effect on tubules stimulated with 10-8 M 5-HT (Fig. 9A). Varying concentrations of
the CAPA analog 2129-SP3[F3]wp-2 were mixed with 10-8 M 5-HT and 10-7 M RhoprCAPA-2.
As previously shown, RhoprCAPA-2 inhibits 5-HT stimulated secretion (Fig. 9B). The analog
prevented the anti-diuretic effect of RhoprCAPA-2 with a statistically significant difference
observed at 10-6 M but failed to return the tubules to its initial rate of 5-HT-stimulated secretion
(Fig. 9B).
41
Table 1: Structure of kinin and CAPA analogs
Analog Structure
Kinin: 2139[Ф1]wp-2 FF[Aib]WGa
CAPA: 2129-SP3[Ф3]wp-2 2Abf-Suc-ATPRIa
42
Fig. 1: A) The effects of injection of 1 µl saline, 1 µl Rhopr-kinin 2 (10-4 M), and 1 µl of the kinin
analog, 2139[Ф1]wp-2 (10-4 M), on the size of blood meal taken by 5th instar R. prolixus. Weight
of insects was measured after 20 minutes of blood-feeding (time 0). The effects of injection of 1
µl of saline and 1 µl of B) Rhopr-kinin 2 (10-4 M) and C) kinin analog 2139[Ф1]wp-2 (10-4 M) on
the rate of diuresis of 5th instar R. prolixus. Weight of insects were measured at time 0 and at 1
hour increments post-feeding for 4 hours. (One-way ANOVA followed by Tukey’s post-hoc test,
slopes tested for significance using an F-test, *=p<0.05. Data are means ± SEM of n=16-20).
43
44
Fig. 2: A) The effects of injection of 1 µl saline, 1 µl RhoprCAPA-2 (10-4 M) and 1 µl CAPA
analog, 2129-SP3[Ф3]wp-2 (10-4 M), on the size of blood meal taken by 5th instar R. prolixus.
Weight of insects was measured after 20 minutes of blood-feeding (time 0). The effects of injection
of 1 µl of saline and 1 µl of B) Rhopr-CAPA 2 (10-4 M) and C) the CAPA analog, 2129-SP3[Ф]wp-
2 (10-4 M), on the rate of diuresis of 5th instar R. prolixus. Weight of insects was measured at time
0 and at 1 hour increments post-feeding for 4 hours. (One-way ANOVA followed by Tukey’s post-
hoc test, slopes tested for significance using an F-test, *=p<0.05. Data are means ± SEM of n=18-
20).
45
46
Fig. 3: Example traces of changes in basal tonus of hindgut contractions in response to A) 10-8 M
Rhopr-kinin 2 and B) 10-8 M kinin analog, 2139[Ф1]wp-2. Downward arrowheads denote
application of peptide, upward arrowheads denote the start of saline wash, and circles denote
vertical deflections due to wash. Dose-response curves displaying changes in basal tonus of
hindgut contractions in response to C) Rhopr-kinin 2 and D) kinin analog, 2139[Ф1]wp-2. (One-
way ANOVA followed by Dunnett’s multiple comparison test, *=p<0.05, **=p<0.01,
***=p<0.001. Data are means ± SEM of n=5).
47
48
Fig. 4: Example traces of changes in basal tonus of hindgut contractions in response to A) 10-8 M
of RhoprCAPA-2 and B) 10-8 M of CAPA analog, 2129-SP3[Ф3]wp-2. Downward arrowheads
denote application of peptide, upward arrowheads denote the start of saline wash, and circles
represent vertical deflections due to wash. C) Dose-response curve displaying changes in basal
tonus of hindgut contractions in response to the CAPA analog, 2129-SP3[Ф3]wp-2. (One-way
ANOVA followed by Dunnett’s multiple comparisons test, *=p<0.05, **=p<0.01. Data are means
± SEM of n=5).
49
50
Fig. 5: A) Example traces of changes in basal tonus of hindgut contractions in response to 10-8 M
Rhopr-kinin 2 and a mixture of 10-8 M Rhopr-kinin 2+10-8 M RhoprCAPA-2. Downward
arrowheads denote application of peptide, upward arrowheads denote the start of saline wash, and
circles denote vertical deflections due to wash. B) 10-8 M RhoprCAPA-2 and C) 10-7 M
RhoprCAPA-2 potentiates the change in basal tonus elicited by varying concentrations of Rhopr-
kinin 2. Change in tension is represented as a percent of maximum tension induced by 10-8 M
Rhopr-kinin 2 on each preparation. (Two-way ANOVA followed by Tukey’s post-hoc test,
*=p<0.05, **=p<0.01. Data are means ± SEM of n=5).
51
52
Fig. 6: The effects of varying concentrations of the CAPA analog, 2129-SP3[Ф3]wp-2, on the
changes in basal tonus elicited by 10-8 M Rhopr-kinin 2. No statistically significant differences
were found. (One-way ANOVA followed by Dunnet’s multiple comparison test, p>0.05. Data are
means ± SEM of n=5).
53
54
Fig. 7: Example traces of changes in the frequency of hindgut contractions in response to A) 10-8
M 5-HT and B) a mixture of 10-8 M 5-HT and 10-7 M of the CAPA analog 2129-SP3[Ф3]wp-2.
Downwards triangles denote vertical deflections of hindgut contractions. C) The effects of varying
concentrations of the CAPA analog 2129-SP3[Ф3]wp-2 on the frequency of hindgut contractions
elicited by 10-8 M 5-HT over a 2-minute period. (One way-ANOVA followed by Dunnett’s
multiple comparison test, *=p<0.05. Data are means ± SEM of n=8).
55
56
Fig. 8: Example traces of changes in the frequency of hindgut contractions in response to A) 10-8
M 5-HT and B) a mixture of 10-8 M 5-HT and 10-6 M RhoprCAPA-2. Downwards triangles denote
contractions. C) The effects of varying concentrations of RhoprCAPA-2 on the frequency of
hindgut contractions elicited by 10-8 M 5-HT over a 2-minute period. (One way-ANOVA followed
by Dunnett’s multiple comparison test, *=p<0.05. Data are means ± SEM of n=8).
57
58
Fig. 9: A) The effects of varying concentrations of the CAPA analog 2129-SP3[Ф3]wp-2 on the
fluid secretion rate of MTs stimulated by 10-8 M 5-HT. No statistically significant differences were
found. B) The antagonist effects of various concentrations of the CAPA analog 2129-SP3[Ф3]wp-
2 on RhoprCAPA-2 (10-7 M) resulted in blocking of the inhibitory effect of Rhopr-CAPA-2 on
MTs stimulated with 10-8 M 5-HT. (One-way ANOVA followed by Dunnet’s multiple comparison
test, *=p<0.05. Data are means ± SEM of n=6).
59
60
Discussion
In R. prolixus, the precisely timed events that occur during post-prandial rapid diuresis are
governed by the neuroendocrine system, therefore investigating the neuroactive chemicals that
may be associated with these specific behaviours and physiology provides insight into how these
processes occur. Also, from an agrochemical perspective, the endocrine system can be used as a
target to disrupt epidemiologically-relevant behaviours in order to prevent disease transmission.
Neuropeptides show great promise in the development of next-generation insecticides, due to their
specificity in terms of function and binding to GPCRs (Audsley & Down, 2015).
As blood feeding is a requirement for the initiation of many developmental and
reproductive processes and the transition to the next instar within R. prolixus, interference with
these events would prove to be quite detrimental (Lange et al., 2016). Insects that were injected
with the kinin analog 2139[F1]wp-2 prior to feeding had a significantly reduced blood meal
compared to saline injected insects which was consistent with previous work where Aib-containing
analogs were found to have antifeedant effects on 5th instar R. prolixus (Lange et al., 2016). In the
hemipteran Acyrthosiphon pisum, while the presence of Aib-containing analogs within the aphid
diets resulted in reduced feeding, and aphicidal activity, 2139[F1]wp-2 failed to reduce aphid
fitness under desiccation and survival stress in M. persicae and M. rosae (Alford et al., 2019b,
Smagghe et al., 2010). These differences may be due to the specificity of the kinin analogs, as
species-specific effects of neuropeptide analogs have been previously observed in D. melanogaster
(Alford et al., 2019a). Injection of the kinin analog did not influence the rate of post-prandial rapid
diuresis which is consistent with the fact that kinins do not play a direct role in post-prandial rapid
diuresis (Lange et al., 2016, Te Brugge et al., 2002; Te Brugge et al., 2009).
61
In contrast to Rhopr-kinins, RhoprCAPA functions as an anti-diuretic hormone through the
inhibition of MT fluid secretion and anterior midgut fluid transport (Paluzzi et al., 2008, Ianowski
et al., 2009). Insects injected with the CAPA analog 2129-SP3[F3]wp-2 prior to feeding took a
significantly increased blood meal compared to saline injected insects and had a significantly
greater rate of post-prandial rapid diuresis within the first hour. As the CAPA analog 2129-
SP3[F3]wp-2 is likely acting upon Rhopr-CAPA receptors as an antagonist (Jiang et al., 2015),
this increased rate of rapid diuresis is likely due to the blocking of the CAPA receptor, thus
preventing the anti-diuretic effects of RhoprCAPA-2. The changes in blood meal size upon
injection of 2129-SP3[F3]wp-2 suggests that RhoprCAPA may also influence feeding. Once an
insect has successfully fed, it has obtained enough nutrients required to moult into the next instar,
and so does not require another blood meal (Buxton, 1930). Since RhoprCAPA-2 is released
towards the end of diuresis, it may also serve as a signal to prevent additional feeding events.
Within R. prolixus, multiple neurohormones such as sulfakinin (Rhopr-SK-1) and Rhopr-CRF/DH
have also been identified as influencing feeding (Al-Alkawi et al., 2017; Mollayeva et al., 2018),
and therefore may co-operatively function in regulating satiety and the motivation to feed. The
injection of RhoprCAPA-2 did not have an impact on the size of blood meal, which suggests that
RhoprCAPA-2 might not influence satiety, but is more a signal to prevent additional feeding
events.
The hindgut of R. prolixus plays an essential role during post-feeding diuresis, as it is
responsible for the excretion of accumulated urine (Maddrell, 1976). In addition, the T. cruzi
parasite is present in the hindgut in its highly infectious stage, and so contraction of the hindgut
results in the release of the parasite along with urine onto the host (Bern et al., 2011). Aib-
containing analogs have previously been shown to be more effective in eliciting myotropic and
62
diuretic effects than their endogenous counterparts in L. maderae and Acheta domesticus
(Nachman et al., 1997; Taneja-Bageshwar et al., 2009). Aib-containing analogs tested on R.
prolixus hindgut were found to be more biologically active than Rhopr-kinins, and in some cases
eliciting irreversible changes in basal tonus (Bhatt et al., 2014). Similar results were obtained with
the kinin analog 2139[F1]wp-2, which caused dose-dependent increases in basal tonus of the
hindgut and was active at concentrations as low as 10-14 M. As described for other Aib-containing
analogs in R. prolixus, the effects of 2139[F1]wp-2 were more difficult to wash off. As these
analogs were synthesized to prevent degradation by endogenous peptidases, the analog may have
a prolonged effect on its target tissue by also influencing the binding to the receptor (Nachman et
al., 2003). These potent changes in physiology induced by the kinin analogs highlight their promise
in future studies for pesticide development.
Despite the presence of CAPA receptors (and indeed pyrokinin receptors) (Paluzzi et al.,
2012; Paluzzi et al., 2008) on the hindgut, none of the three CAPA neuropeptides (RhoprCAPA-
1, RhoprCAPA-2, or Rhopr-pk1) were found to have any direct effect on hindgut contractions.
However, the CAPA analog 2129-SP3[F3]wp-2 induced hindgut contraction in a dose-dependent
manner, but its effects were not as intense as Rhopr-kinin 2 or the kinin analog 2139[F1]wp-2.
Interestingly though, RhoprCAPA-2 potentiated the effect of Rhopr-kinin 2. Since RhoprCAPA-
2 is released as a signal to terminate diuresis, it may also assist the hindgut in excretion of the
remaining urine. This potentiation effect was not observed with RhoprCAPA-1 or Rhopr-pk1, nor
was it seen with the CAPA analog 2129-SP3[F3]wp-2.
63
The potentiation effect of RhoprCAPA-2 may be due to the interaction of separate second
messenger pathways after GPCR activation. As the kinin receptor has not yet been characterized
within R. prolixus, the associated second messenger pathway is currently unknown. Within other
insects, the kinin receptor has been associated with an increase in intracellular Ca2+ via the inositol
phosphate (IP3) pathway. Activation of the IP3 pathway increases IP3 levels within the cytoplasm
which later binds to an IP3-sensitive Ca2+ channel on the endoplasmic reticulum. Following
release, Ca2+ induces muscle contraction by enabling cross-bridge cycling (Radford et al., 2002;
Tehrzaz, et al., 1999; Beyenbach, 2003; Pietrantonio et al., 2005; Kuo & Ehrlich, 2015). In R.
prolixus, activation of the CAPA receptor in the MTs results in an increase in cGMP, which in
turn activates a phosphodiesterase that degrades cAMP, thereby lowering cAMP levels. (Paluzzi
& Orchard, 2006; Paluzzi et al., 2013). Here, we propose that the CAPA receptor may be
constitutively active within the hindgut, with cGMP stably keeping cAMP levels low. Upon
activation of both the kinin and CAPA receptors, the increase in cGMP may participate in the IP3
pathway to increase intracellular Ca2+ resulting in stronger contractions. In vertebrates, cGMP
signaling is implicated in both stimulating or inhibiting contraction (Fischmeister et al., 2005;
Fellner & Arendshorst, 2009). Within vertebrate cardiac muscle, cGMP has shown to have
excitatory effects through the stimulation of Ca2+ channels, resulting in an increase of intracellular
Ca2+. These Ca2+ channels are responsible for the excitation-contraction coupling within the muscle
(Wang et al., 1999; Fischmeister et al., 2005). In contrast are the stimulatory effects of the CAPA
analog 2129-SP3[F3]wp-2. This analog appears to be an antagonist of the CAPA receptor and so
blocks the cGMP-mediated cAMP degradation via activation of a phosphodiesterase, in turn
allowing cAMP to increase and stimulate hindgut contractions. In order to further investigate the
possible mechanism by which this kinin/CAPA interaction occurs, the effects of RhoprCAPA-2
and 2129-SP3[F3]wp-2 were assessed with 5-HT. 5-HT has myostimulatory effects on the
64
hindgut, via an increase in cAMP levels (Orchard, 2006). A potentiation effect was observed with
co-application of 5-HT and 2129-SP3[F3]wp-2 on the frequency of hindgut contractions. This is
likely due to the antagonist action on the CAPA receptors, preventing the RhoprCAPA-mediated
cAMP decrease. Conversely, the effects of 5-HT were inhibited by RhoprCAPA-2, due to the
increase in cGMP, activation of a phosphodiesterase, and degradation of cAMP.
The MTs are critical in allowing R. prolixus to return to a homeostatic state following a
blood meal. As diuretic hormones such as 5-HT, Rhopr-DH31, and Rhopr-CRF/DH function to
stimulate secretion within the tubules, Rhopr-CAPA 2 is required as a signal to abolish this
secretion to prevent excess ion and water loss (Paluzzi et al., 2008; Orchard, 2006). The CAPA
analog 2129-SP3[F3]wp-2 was found to interfere with Rhopr-CAPA 2’s ability to inhibit 5-HT-
stimulated MT secretion, that is to say, it is an antagonist of the CAPA receptor. Within D. suzukii,
2129-SP3[F3]wp-2 had a protective effect as females injected with the analog had a significantly
increased survival rate. Within D. melanogaster, CAPA functions as a diuretic hormone (Kean et
al., 2002), with desiccation survival linked to the regulation of fluid secretion (Terhzaz et al.,
2015). This supports the function of 2129-SP3[F3]wp-2 as a CAPA receptor antagonist, as it
prevents CAPA-stimulated fluid secretion in D. melanogaster, thereby increasing survivability
under desiccation stress (Alford et al., 2019a). Within M. persicae and M. rosae, however,
injection of 2129-SP3[F3]wp-2 resulted in accelerated mortality under desiccation and starvation
stress (Alford et al., 2019b). These effects of the CAPA analog may be due to the lack of MTs
within aphids, therefore CAPA may not play a direct role in desiccation (Jing et al., 2015).
In summary, these results display the efficacy in which the biostable neuropeptide analogs
are able to induce potent changes in physiology, thus showing potential use in the development of
65
pest control strategies. The need for highly coordinated release of neuroactive chemicals within R.
prolixus is a necessity in order for the insect to successfully gorge on blood. Given the bugs
susceptibility to predators in its engorged state, rapid elimination of water and salt is required
(Orchard, 2006). These analogs are able to successfully disrupt this coordination. The novel
interaction between Rhopr-kinin 2 and RhoprCAPA-2 further highlights the importance of
coordinated release of these neuropeptides throughout the life cycle of R. prolixus. Investigating
the intracellular mechanisms by which the actions of the neuroactive chemicals function provides
insight into the mode of action of epidemiologically-relevant behaviours, which can also aid in the
development of next-generation pest control strategies.
Acknowledgements
The authors would like to thank Stuti Joshi for maintaining the colony and Allison Strey
(ARS-USDA) for technical assistance. This work was financially supported by Natural Sciences
and Engineering Research Council of Canada Discovery Grant [RGPIN 2014-06253 to AL, and
RGPIN 8522-12 to IO].
66
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Chapter 3 Identification and Cloning of the Kinin Receptor in the Chagas
Disease Vector, Rhodnius Prolixus
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Abstract
Within invertebrates, the kinin family of neuropeptides is responsible for the modulation
of a host of physiological and behavioural processes. In Rhodnius prolixus, kinins are primarily
responsible for eliciting myotropic effects on various feeding and diuresis-related tissues. Kinins
function through binding to G-protein coupled receptors (GPCRs), likely utilizing intracellular
Ca2+ as a second messenger. Here, the R. prolixus kinin receptor (RhoprKR) has been successfully
isolated, cloned and sequenced from the central nervous system (CNS) and hindgut of R. prolixus.
Sequence analysis show high similarity between RhoprKR with other cloned invertebrate kinin
receptors, along with the presence of various highly conserved residues. The expression profile of
RhoprKR shows expression of the RhoprKR transcript throughout the R. prolixus gut, suggesting
a role of Rhopr-kinins in various aspects of feeding and digestion. RNA interference (RNAi)-
mediated knockdown of the RhoprKR transcript resulted in a significant reduction of hindgut
contractions in response to Rhopr-kinin 2 and an Aib-containing kinin analog. dsRhoprKR-
injected insects also consumed a significantly larger blood meal, suggesting a role of Rhopr-kinins
in satiety. Overall, these findings highlight the role of the kinin signaling system in R. prolixus,
and the effectiveness of RhoprKR as a target in RNAi-mediated pest control strategies.
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Introduction
Neuropeptides comprise the largest family of neuroactive chemicals and are involved in
many essential physiological and behavioural processes within insects. The effects of these
neuropeptides are directed through activation of G protein-coupled receptors (GPCRs), the largest
group of membrane receptors (Altstein & Nässel, 2010; Lismaa & Shine, 1992). In insects, studies
have examined the potential use of these neuropeptides in pest control strategies, since they are
more environmentally-friendly than traditional pesticides and can elicit effects at low
concentrations (Nachman & Smagghe, 2011; Gäde & Goldsworthy, 2003). In the context of
pharmaceutical development, GPCRs are often used as targets for novel drugs, with 40-50% of
drugs acting upon GPCRs (Garland, 2013). However, GPCRs have only recently been targeted in
pest control strategies within insects, despite their potential use in the development of next-
generation pesticides (Audlsey & Down, 2015). Increasing our understanding of neuropeptide
families and their GPCRs can provide great insight into how crucial processes can be disrupted,
preventing the spread of relevant pest species and disease vectors (Audsley & Down, 2015;
Nachman & Smagghe, 2011; Nachman, 2009).
In the Chagas disease vector, Rhodnius prolixus, post-feeding physiology has been widely
studied and is under regulation by various neuroendocrine factors (Orchard, 2006; Coast et al.,
2002; Orchard, 2009). Diuresis and excretion are considered to be epidemiologically-relevant
behaviours, as they are implicated in the transmission of Chagas disease (Orchard, 2006; Orchard
& Paluzzi, 2009). Excretion occurs through contractions of the hindgut, where the Trypanosoma
cruzi parasite exists in the highly infectious stage of its life cycle (Bern et al., 2011). Contractions
of the hindgut result in the transmission of the parasite onto the host, and therefore possible
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infection (Bern et al., 2011). From a medical perspective, neuropeptides that modulate hindgut
contractions are compounds of interest, as they can be targeted in an effort to disrupt excretion.
The kinins, first identified for their myotropic effects in the cockroach Leucophaea
maderae, are a family of neuropeptides with the following conserved C-terminal pentapeptide
sequence: FX1X2WG-amide, where X1 can be Ser, Phe, His, Asn, or Tyr, and X2 can be Ser, Pro
or Ala (Holman et al., 1986a, b; Holman et al., 1987a, b). This core pentapeptide sequence is
required for the neuropeptide to be biologically active (Coast et al., 1990; Nachman et al., 1991).
Kinins have now been identified in a number of species and are implicated in a diverse set of
functions (Rosay et al., 1997; O’Donnell et al., 1998; Tehrzaz et al., 1999; Kim et al., 2006; Bhatt
et al., 2014; Zandawala et al., 2018). The R. prolixus kinin (Rhopr-kinin) transcript encodes for
the most kinins found in any species, with eighteen predicted kinins and precursor associated
species (Te Brugge et al., 2011; Bhatt et al., 2014). Within R. prolixus, Rhopr-kinins are primarily
known for their myotropic effects, eliciting contractions of the hindgut, midgut, and salivary glands
(Orchard & Te Brugge, 2002; Te Brugge et al., 2009; Te Brugge et al., 2011; Bhatt et al., 2014).
Given the ability of Rhopr-kinins to stimulate hindgut contractions, they are a neuropeptide of
interest, as they are directly implicated in the transmission of the T. cruzi parasite, and Chagas
disease (Bern et al., 2011).
Kinin GPCRs belong to the family A GPCRs, which is the largest and most studied family
of GPCRs, with the first kinin receptor characterized in the snail, Lymnaea stagnalis (Cox et al.,
1997; Munk et al., 2016). Following this, kinin receptors have been characterized in a small
number of insects including Drosophila melanogaster, Aedes aegypti, and Anopheles stephensi
(Radford et al., 2002; Radford et al., 2004; Pietrantonio et al., 2005). Kinin receptors may function
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by increasing intracellular Ca2+ through the inositol phosphate (IP3) pathway, as activation of this
pathway was observed within the Malpighian tubules (MTs) of D. melanogaster and A. aegypti
(Cady and Hagedorn, 1999; Radford et al., 2002; Pietrantonio et al., 2005). Interestingly, the kinin
receptor within A. aegypti functions as a multiligand receptor for the three Aedes kinins, with the
three kinins utilizing different intracellular pathways (Pietrantonio et al., 2005). Currently, the
kinin receptor within R. prolixus has not yet been characterized, but analysis of potential GPCRs
within the R. prolixus transcriptome has identified a possible kinin receptor candidate
(RPRC000494) (Ons et al., 2016).
Advances in molecular biology have allowed the development of various tools that can be
used in the creation of novel strategies to combat pests and disease vectors. RNA interference
(RNAi) is a common research tool in insects, and can be used in pest control development, since
it can specifically target species (Vogel et al., 2019). RNAi has been successful at inducing
phenotypic changes that decrease overall fitness, such as reduction of cardiac output through
silencing of the crustacean cardioactive peptide (CCAP) transcript in A. gambiae (Estévez-Lao et
al., 2013) and reduced fecundity after silencing of the kinin receptor in Rhipicephalus microplus
(Brock et al., 2019). Within R. prolixus, RNAi has been utilized in a similar manner, with silencing
of the CCAP receptor leading to a decrease in basal heartbeat frequency and disrupting ecdysis
(Lee et al., 2013). In addition, R. prolixus injected with chitin synthase (CHS) double-stranded
RNA (dsRNA) had severe cuticle deformations which interfered with mobility and longevity, and
adult females injected with CHS dsRNA had overall reduced oviposition (Mansur et al., 2014).
In the case of larger insects such as R. prolixus, dsRNA is often delivered through injection
which can induce phenotypic changes that may not be associated with the intended knockdown
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target, or cause trauma due to the injection wound (Whitten et al., 2016). In Acyrthosiphon pisum,
injected of dsRNA resulted in unexpected changes in gene expression and disruptive effects
resulting in mortality (Jauber-Possamai et al., 2007). A novel form of RNAi delivery, known as
symbiont-mediated RNAi aims to overcome the limitations of large-scale dsRNA delivery. This
form of delivery involves the integration of dsRNA into the symbiotic bacteria of a target species,
thus evoking RNAi within the host (Whitten et al., 2016). Symbiont-mediated RNAi has been
successfully tested on R. prolixus, inducing various knockdown phenotypes. As these recombinant
bacteria can be delivered through a blood meal or through coprophagy within R. prolixus
populations, large-scale application of this RNAi strategy can prove to be quite feasible (Whitten
et al., 2016).
In this study, the R. prolixus kinin receptor (RhoprKR) was isolated and cloned. The
expression profile of the RhoprKR transcript within the tissues of 5th instar R. prolixus using
quantitative PCR (qPCR) was determined. As kinins are responsible for the contraction of hindgut
muscle within R. prolixus (Bhatt et al., 2014; Te Brugge et al., 2011), RNAi was utilized to
knockdown the RhoprKR transcript followed by hindgut contraction assays to confirm
knockdown. In addition, the effects of RhoprKR knockdown was assessed on in vivo feeding and
diuresis. Identification of RhoprKR will assist in the development of next-generation pest control
strategies where this receptor can be targeted to disrupt the overall physiology of R. prolixus and
therefore release of the parasite onto the host.
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Materials and Methods
Animals
5th instar male and female R. prolixus were obtained from a colony at the University of
Toronto Mississauga. Insects were reared in incubators at 25oC and 50% humidity and were fed
defibrinated rabbit blood (Cedarlane Laboratories, Burlington, ON, Canada) once in each instar.
Chemicals
Rhopr-kinin 2 was custom synthesized by Genscript (Piscataway, NJ, USA). The
neuropeptides were then reconstituted in double-distilled water in stock solutions at 10-3 M, and
later stored at -20oC. The Aib-containing insect kinin analog (2139[F1]wp-2) with the following
amino acid sequence Phe-Phe-Aib-Trp-Gly-NH2 (Nachman et. al., 1996) was prepared in 80%
aqueous acetonitrile (ACN) at a concentration of 10-3 M and stored at 4oC. Rhopr-kinin 2 and
2139[F1]wp-2 were later diluted in physiological saline (150 mM NaCl, 8.6 mM KCl, 2.0 mM
CaCl2, 4.0 mM NaHCO3, 8.5 MgCl2, 0.02 mM HEPES and 34 mM glucose in pH 7.0) to various
concentrations for use in bioassays.
Identification and Cloning of cDNA Sequences Encoding the R. prolixus Kinin Receptor
The genome, transcriptome, and peptidome of R. prolixus was obtained from
vectorbase.org and uploaded into Geneious 8.1 (Auckland, New Zealand). A BLAST search was
performed against the R. prolixus transcriptome using the D. melanogaster kinin receptor
(Q9VRM0) and A. aegypti kinin receptor (Q5EY37) as templates, giving hits for an annotated
transcript sequence (RPRC00494), which has been previously predicted to be a Rhopr-kinin
receptor (Ons et al., 2016). Following this, a nucleotide sequence alignment was performed with
the D. melanogaster and A. aegypti kinin receptors, and highly conserved cDNA sequences within
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the predicted Rhopr-kinin receptor (RhoprKR) were amplified by gene-specific primers (Table
S1). OneTaq ® DNA Polymerase (NEB, Whitby, ON, Canada) was used for all PCRs, and
reactions were performed using Bio-Rad’s s100 thermocycler (Bio-Rad Laboratories, Mississauga,
ON, Canada). PCR products were extracted from 1.2% agarose gel using an EZ-10 Spin Column
DNA Gel Extraction Kit (Bio Basic, Markham, ON, Canada) and later cloned using the pGEM-T
Easy Vector (Promega, Madison, WI, USA). Bacteria that had successfully taken up the insert
were inoculated and left to grow overnight. Using the EZ-10 Spin Column Plasmid DNA MiniPrep
Kit (Bio Basic, Markham, ON, Canada), inserts were extracted, and sent for Sanger sequencing at
Macrogen USA (Macrogen, Brooklyn, NY, USA).
To successfully amplify the 5’ and 3’ regions of RhoprKR, 5’ and 3’ RACE was performed
using the SMARTer® RACE 5’/3’ Kit (Takara Bio USA, Mountain View, CA, USA). The CNS
and hindgut was dissected and placed in nuclease-free phosphate-buffered saline (PBS) (Sigma
Aldrich, Oakville, ON, Canada). Following this, total RNA extraction was performed using the
EZ-10 Spin Column Total RNA Miniprep Super kit (Bio Basic, Markham, ON, Canada). cDNA
was synthesized using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems,
Mississauga, ON, Canada) to be later used to synthesize 5’ and 3’ RACE-ready cDNA. Gene
specific forward and reverse primers were developed for the 5’ and 3’ end of the RhoprKR
transcript, to be amplified using the 5’ and 3’ RACE-ready cDNA (Table S2). Using the initial 5’
and 3’ RACE products as DNA templates, a nested PCR was performed with the additional
downstream 5’ and 3’ primers to ensure specificity of the amplified sequence. 5’ and 3’ RACE
reactions were performed using the SMARTer® RACE 5’/3’ Kit protocol, and products of the
RACE reactions were later purified using the High Pure PCR Product Purification Kit (Roche
Applied Science, Penzberg, Germany), cloned, and sequenced as described above.
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Sequence Analysis
Following sequencing of RhoprKR, the structural and biochemical features of the receptor
were analyzed using various online tools. The seven transmembrane domains for RhoprKR were
predicted by TMHMM Server v.2.0 (http://www.cbs.dtu.dk/services/TMHMM/). The potential N-
linked glycosylation sites were predicted using the NetNGlyc 1.0 Server
(http://www.cbs.dtu.dk/services/NetNGlyc/) with potential phosphorylation sites predicted using
the NetPhos 3.1 Server http://www.cbs.dtu.dk/services/NetPhos/). Lipid modification sites for
palmitoylation of cysteine residues were predicted by GPS-Lipid
(http://lipid.biocuckoo.org/webserver.php). To predict the exon-intron boundaries within the
sequence, BLAST was performed, following confirmation with a fruit fly splice site prediction
tool. (http://www.fruitfly.org/seq_tools/splice.html). Alignment of RhoprKR was performed with
cloned kinin receptors of invertebrate species and other R. prolixus receptors using the MUSCLE
alignment tool (https://www.ebi.ac.uk/Tools/msa/muscle/).
Spatial Expression of Rhopr-kinin Receptor in Feeding and Diuresis-Related Tissues
Expression of RhoprKR was examined in 5th instar R. prolixus, 3-5 weeks post-feeding as
4th instars. The CNS, fat body (FB), dorsal vessel (DV), salivary glands (SG), foregut (FG),
posterior midgut (PMG), anterior midgut (AMG), hindgut (HG), and MTs were dissected using
nuclease-free PBS. Tissues were dissected and pooled into three different biological replicates
(n=3). Total RNA extraction was performed using an EZ-10 Spin Column Total RNA Miniprep
Super kit with cDNA synthesized using a High-Capacity cDNA Reverse Transcription Kit. cDNA
was used for quantitative PCR (qPCR) reactions. cDNA was diluted by 10-fold and amplified
using qPCR primers for RhoprKR (Table S4) and reference genes (α-tubulin, β-actin, ribosomal
protein 49) (Table S1). qPCR reactions were performed using Bio-Rad’s CFX96 TouchTM Real-
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Time PCR Detection System (Bio-Rad Laboratories, Mississauga, ON, Canada). Three technical
replicates were performed per tissue, with a non-template control for each biological replicate.
Expression of the transcript levels in each tissue were calculated relative to an average of the
reference genes using the ∆Ct method.
Double Stranded RNA Synthesis
Gene-specific primers were developed for a partial cDNA sequence from the RhoprKR
transcript and the ampicillin resistance gene (ARG) (to be used as a control). These primers were
also conjugated with 23 bases of the T7 RNA polymerase promoter on the 5’ end (Table S3). The
cDNA sequence for RhoprKR was amplified using R. prolixus CNS cDNA, with ARG amplified
from the pGEM-T Easy Vector system, both via PCR. These PCR products were purified using
the High Pure PCR Product Purification Kit, to be used as a DNA template for the following T7
PCR reactions. The sequences possessing the T7 promoters were amplified via PCR, and later
purified with the High Pure PCR Product Purification Kit. Double stranded RNA (dsRNA) was
synthesized using the purified products of the T7 reactions using the T7 RiboMAX Express RNAi
System (Promega, Madison, WI, USA). Following synthesis, the dsRNA was precipitated with
isopropanol, eluted in InvitrogenTM UltraPureTM DNase/RNase-Free Distilled Water (Thermo
Fisher Scientific, Waltham, MA, USA) then quantified using a nanodrop at 260nm wavelength to
assess its concentration and quality. The dsRNA was later resuspended in InvitrogenTM
UltraPureTM DNase/RNase-Free Distilled Water at a final concentration of 2 μg/ml.
dsRNA Delivery
5th instar R. prolixus were injected through the membrane at the junction of the hind leg
with the abdomen with 1 μl of 2 μg/ml RhoprKR dsRNA (dsRhoprKR) or ARG dsRNA (dsARG)
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using a 10 μl Hamilton syringe (Hamilton Company, Reno, Nevada, USA). Following injection,
R. prolixus were left to recover at room temperature for two hours, then placed in incubators at
25oC and 50% humidity.
Knockdown Verification using Quantitative PCR
Three pools of CNS and hindgut tissue (n=3) were dissected using nuclease-free PBS from
R. prolixus at 2 days and 7 days post-injection with dsRhoprKR or dsARG, and RNA extraction
was performed with the EZ-10 Spin Column Total RNA Miniprep Super kit. cDNA was
synthesized using the High-Capacity cDNA Reverse Transcription Kit and diluted 10-fold to be
used for qPCR. To verify the knockdown efficiency, qPCR was performed as described above,
with changes in transcription levels measured using the 2-∆∆Ct method. Insects from day 7 post-
injection (highest knockdown efficiency) were selected to be used for hindgut contraction assays
and feeding bioassays.
Hindgut Contraction Assay via Force Transducer
The R. prolixus hindgut was isolated under physiological saline along with the cuticle at
the posterior end and fixed onto a Sylgard-coated dish using minutien pins through the cuticle. The
isolated hindgut was bathed in 200 μl of physiological saline. One end of a fine silk thread was
tied to the anterior end of the hindgut with the other end tied to a Grass FT03 force transducer
(Astro-Nova Inc., Rhode Island, USA). Using a Picoscope 2204 Oscilloscope (Pico Technology,
Cambridgeshire, UK), the amplitude of basal tonus changes was recorded. Tissues were first
equilibrated in physiological saline for 10 minutes followed by application of Rhopr-kinin 2 or the
kinin analog 2139[F1]wp-2. 100 μl of various concentrations of Rhopr-kinin 2 or the kinin analog
were applied to the bath concurrent with removal of 100 μl of saline to ensure the bath volume
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remained constant. In between doses the preparations were washed with saline and the bath volume
was maintained at 200 μl.
Feeding Bioassay
On day 7 post-injection, dsARG-injected and dsRhoprKR-injected insects were weighed
before feeding, then placed in a 10 cm diameter glass jar and fed on 20 mL of a warm saline
consisting of 0.15 M NaCl and 10-3 M ATP (Lange et al., 1988; Friend, 1965) for 20 minutes.
Following feeding, insects were individually weighed immediately after feeding (time 0), and then
placed in individual cubicles. To measure the rate of diuresis, the weights of individual insects
were then recorded at 1,2,3, and 4-hour time points. Insects that fed less than 1 times their initial
body weight were excluded as they were not considered to have fed successfully. Insects that were
also punctured during the weighing process were omitted from the data.
Results
Structure and Sequence Analysis of RhoprKR
Following an in silico analysis of potential kinin GPCRs within the R. prolixus
transcriptome, partial sequences of the candidate kinin receptor were amplified using PCR. 5’ and
3’ RACE was then utilized to amplify the open reading frame (ORF) and the 5’ and 3’ untranslated
regions (UTR). The length of the ORF of RhoprKR is 1285 bp, which translates to 415 amino
acids (Fig. 1A). The ORF spans 8 exons, separated by 7 introns (Fig. 1B). The 5’ UTR is 40 bp,
and the 3’ UTR is 387 bp, with two stop codons observed in the 5’ UTR (Fig. 1A). Following
synthesis, GPCRs undergo post-translational modifications to ensure complete functionality which
include N-glycosylation, palmitoylation of Cys residues, and phosphorylation (Kristiansen, 2004;
Duvernay et al., 2005). Within the RhoprKR transcript, N-glycosylation is predicted to occur at
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Asn2, and Asn25 (Fig. 1A). Phosphorylation sites are predicted on 4 Thr residues, 2 Tyr residues,
and 15 Ser residues, with palmitoylation predicted to occur on Cys338 (Fig. 1A).
Alignment of cloned GPCRs of the kinin family with other R. prolixus receptors reveals
considerable identity and conserved features across the kinin receptors but not against GPCRs for
other peptide families (Fig 2), including other family A and family B receptors (Lee et al., 2016).
Amongst the aligned kinin receptors there is approximately 42% pairwise identity across the ORFs.
Sequence similarity is high within TM1, TM2, TM3, TM6, and TM7, with TM4 and TM5 showing
less similarity amongst the receptors (Table 1). Family A GPCRs are characterized by a DRY (and
in some cases DRH and ERY) motif, and an NPxxY domain (Holmes et al., 2000; Capra et al.,
2004; Radford et al., 2004; Pietrantonio et al., 2005; Munk et al., 2016). This motif is seen in all
of the aligned family A receptors, while it exists as DRY/DRH in the kinin receptors, but as ERY
in the sulfakinin receptors, Rhopr-SKR-1 and Rhopr-SKR-2, and in the CAPA receptor, capa-r1
(Fig. 2). The NPxxY domain is also observed within all of the family A receptors but exists as
NPFIY in the kinin receptors (Fig. 2). Conserved Cys residues present in the first and second
extracellular loops, predicted to form a disulphide bridge, are conserved in all of the kinin receptors
except for L. stagnalis (Holmes et al., 2000; Radford et al., 2004; Pietrantonio et al., 2005) (Fig.
2). Unlike the TMs, a high degree of similarity is not observed within the N-terminus and C-
terminus. However, some conserved Ser and Thr residues within the C-terminus may represent
conserved phosphorylation sites across the receptors (Radford et al., 2004).
Spatial Expression of RhoprKR
Transcript expression level of RhoprKR was measured in the CNS and various tissues of
5th instar R. prolixus using qPCR. High expression of the RhoprKR transcript is observed in the
84
CNS and HG (Fig. 3). There are similar transcript expression levels within the FG and PMG, with
slightly lower expression in the AMG (Fig 3). Expression of the RhoprKR transcript is also
observed in the SG, but expression levels are lower than those seen in the gut (Fig. 3). Low
expression is observed in the FB, DV, MTs (Fig 3).
Knockdown of RhoprKR Transcript
To verify the likelihood that this RhoprKR transcript is a kinin GPCR, RNAi was utilized
to reduce the transcript expression within 5th instar R. prolixus. Changes in RhoprKR transcript
expression was verified in the CNS and hindgut using qPCR, with a reduction in transcript
expression of 13% observed in the CNS, and 18% in the hindgut on day 2 post-dsRNA injection.
On day 7 post-dsRNA injection, a 76% reduction in transcript expression was observed in the CNS
and a 70% reduction in transcript expression in the hindgut (Supp. Fig.1).
Effects of RhoprKR knockdown on Hindgut Contractions
Following RNAi-mediated knockdown of the RhoprKR transcript in R. prolixus, hindgut
contraction assays were performed on insects 7 days-post injection. Both Rhopr-kinin 2 and the
kinin analog 2139[F1]wp-2 elicited smaller changes in basal tonus of hindgut contraction from
dsRhoprKR-injected insects than dsARG-injected insects (Fig. 4). Relative to dsARG,
dsRhoprKR-injected insects had a 43 % reduction in response to 10-8 M Rhopr-kinin 2, with a 54
% reduction in response to a dose of 10-8 M 2139[F1]wp-2 (Fig 4).
Effects of RhoprKR Knockdown on in vivo Feeding and Diuresis
To observe the effects of RhoprKR knockdown on feeding and diuresis, the blood meal
size and excretion rate for dsARG-injected and dsRhoprKR-injected insects was examined. In
85
comparison to dsARG-injected insects, dsRhoprKR-injected insects had consumed a significantly
larger meal, feeding an average of 12 times their initial body weight (Fig 5A). In addition, a higher
number of dsRhoprKR-injected insects fed on a meal much greater than their body weight, with
17 insects feeding over 10 times their body weight. For dsARG-injected insects, 9 insects fed over
10 times their body weight, while 7 insects fed 2-4 times their body weight (Fig. 5B). As post-
feeding diuresis occurs over a time period of 3-4 hours following feeding, the weight loss due to
excretion was monitored for the dsARG-injected and dsRhoprKR-injected insects to examine
changes in diuresis rate. Despite a difference in artificial blood meal size, there was no difference
in the rate of diuresis over the measured 4 hours (Fig 5B)
86
Fig. 1: A) The cDNA sequence of RhoprKR with the predicted amino acid sequence. Nucleotide
number is on the right of the sequence, with amino acid number bolded below the nucleotide
numbers. Predicted transmembrane domains within the sequence are highlighted in grey. Stop
codons in the 5’UTR are boxed in green. Predicted N-glycosylation sites are shaded light blue and
underlined, potential phosphorylation sites boxed in black, and a potential palmitoylation of Cys
residue in green. B) The exon map of RhoprKR, with the open reading frame (ORF) and
exons/introns. The ORF of RhoprKR is denoted by the solid black box spanning the exons, with
numbers above exons representing exon length and numbers below introns representing intron
length.
87
B)
35
A)
105 194 150 126 217 102 92 654 bp
bp107098 19390 14682 1348 177 851 105 187
atg aac tgc agt ttt cta gaa gac gaa cta ggg cct tta cca cca agc gca aat tgt tca 100
M N C S F L E D E L G P L P P S A N C S 20
tgg ctc ctt cac aat caa tca gtg tat ttc tat gaa gaa agt ctg tac gaa gtt cct gcc 160
W L L H N Q S V Y F Y E E S L Y E V P A 40
ggt gta ata gta tta cta tca gta ttc tat ggc acg ata tct gtg gtg gca gtt ggc gga 220
G V I V L L S V F Y G T I S V V A V G G 60
aat ttt ctg gtt atg tgg att gtg gcc act agt aga cgc atg cag aat gtt act aac tgt 280
N F L V M W I V A T S R R M Q N V T N C 80
ttt ata gca aat cta gcc ttg gct gat ata gtt atc ggc cta ttc gct ata cca ttt caa 340
F I A N L A L A D I V I G L F A I P F Q 100
ttc caa gca gca ctg ttg cag agg tgg aac ctg cca aat ttt atg tgc cct ttt tgc cca 400
F Q A A L L Q R W N L P N F M C P F C P 120
ttc gtc cag gtg ctc agc gtc aat gtt agt gta ttc aca ctg acg gcc att gct gta gat 460
F V Q V L S V N V S V F T L T A I A V D 140
aga cat cgg gca gtc ctt aac cca tta agt gct cca ccc tca aaa tta aga gct aaa gct 520
R H R A V L N P L S A P P S K L R A K A 160
tta ttg ggc gct att tgg att ttg gct gct atc ctt gca aca cca atg gct gta gca tta 580
L L G A I W I L A A I L A T P M A V A L 180
aat gta act tat gtt gaa gaa aat gat cat gtt ggt cat gtt tac acc aaa cca ttt tgt 640
N V T Y V E E N D H V G H V Y T K P F C 200
ata aat aca aaa ctg tca aat aac cat atg atg gcc tac agg atg ata ttg gta tca gtc 700
I N T K L S N N H M M A Y R M I L V S V 22
caa tat ctg aca ccg ttg tgt gtc ata tca tat gcc tat gca aaa atg gca ctc agg ttg760
Q Y L T P L C V I S Y A Y A K M A L R L 240
tgg gga tca aga gct cct ggt aat gct caa cat tct agg gat gca aat tta atg aga aat 820
W G S R A P G N A Q H S R D A N L M R N 260
aag aaa aag gta atc aaa atg tta gtt ata gtg gtg gct cta ttt gca ata tgt tgg ctt 880
K K K V I K M L V I V V A L F A I C W L 280
cca tta caa act tac aat gtt tta caa gac ata ttt cca caa att aat ggg tac aga tat 940
P L Q T Y N V L Q D I F P Q I N G Y R Y 300
att aat ata att tgg ttt tgt tgt gat tgg ctg gca atg agt aat tcc tgt tac aat ccg 1000
I N I I W F C C D W L A M S N S C Y N P 320
ttt ata tac ggt att tac aat gaa aag ttt aag caa gaa ttt caa caa cga tgc ccg ttc 1060
F I Y G I Y N E K F K Q E F Q Q R C P F 340
agt aga aga aga aag tgg act cat ggt ttt ggt gcc ggt ggt agt gac agt tta gat cta 1120
S R R R K W T H G F G A G G S D S L D L 360
gat aaa aca ata cat cgt ttt ggt agt gtt aac cgt aat tcg tct cgc tgg ata aga tac 1180
D K T I H R F G S V N R N S S R W I R Y 380
tca tcc cgt gtg caa tat act cca gca caa cat tac att tac cac tgt gcc aat tcc aat 1240
S S R V Q Y T P A Q H Y I Y H C A N S N 400
act gta cac cac agt tcc caa tca gaa ata gaa gag ctt tgt ctg taa 1288
T V H H S S Q S E I E E L C L - 416
5’-tactatctgtcatgtgtaatacgtcatctaactaaagatt 40
atactattattacaattaacttgtgaacatcttgcttcagaaatattatgcagtgtaacaataacttttggacattttttatgaaccaaaatttttacaaatatctgtagatttagtgacgaaattattgtcggaattcacaattgtattttcacattttcaattggaattctaaaatgaaatg
aattaggtattcgtggtacgtttccttatattttcctgcttctgaagttaggtgtgcataccctcgaaaatcattaacaagtgtgatgtattaccctaaagagctagaaaatctctaatggacagcatttaactgaatataatttttggggcacatttaaataaaaccatttatctcaaatgagtcgctgtacaaatgtaaaa-3’
16751675
88
Fig. 2: Multiple sequence alignment of cloned invertebrate kinin receptors and various cloned R.
prolixus receptors. Sequences shaded in black represent identical amino acids across sequences
and 60% similar sequences are shaded in grey. Bolded red lines above sequences represent
predicted RhoprKR transmembrane domains. The conserved DRY/DRH motif, NPxxY domain,
and conserved Cys residues are boxed in red. Abbreviations: Rhopr-CRF/DH-R2: R. prolixus
CRF/DH receptor 2 (A0A191UP91); L. stagnalis: Lymnaea stagnalis kinin receptor (P92045); D.
melanogaster: Drosophila melanogaster kinin receptor (Q9VRM0); A. stephensi: Anopheles
stephensi kinin receptor (Q69V6); A. aegypti: Aedes aegypti kinin receptor (Q5EY37); R.
microplus: Rhipicephalus microplus kinin receptor (Q9NHA4); RhoprCAPA-R1: R. prolixus
CAPA receptor variant A (D6P3E5): Rhopr-SKR-1: R. prolixus sullfakinin receptor 1
(MK513659); SKR-2: R. prolixus sullfakinin receptor 2 (MK513660).
89
Rhopr-CRF/DH-R2 1 -----GLLCWPNTPPGVTAYLPCV-------AEIDNVKYDTNQNASRICYENGTWANQTD L. stagnalis 1 MSQIESMSEQA---AVI--FIEQANQDLDNVSGNDVSSFFYNET--------TTLF---- D. melanogaster 1 -------------------------------MAMDLIEQE--SR--------LE-FLP-- A. stephensi 1 MQ------------------------------ATDITAYHTAYN--------YTLNQS-- A. aegypti 1 MR------------------------------AVDGIAFHYANN--------NTLNGS-- R. microplus 1 MTSLPGMTLDPSAPPPL--LLDSS------YVSPDYGN--------------LSLLSS-- RhoprKR 1 --------------------MNCS------FLEDELGPLPPSAN--------CSWLLH-- RhoprCAPA-R1 1 ------------------------------MNSFDIIETVT-NS--------TPVNVS-- Rhopr-SKR-1 1 ------------------------------------------------------------ Rhopr-SKR-2 1 ---------------------------------------MR-NN--------TEATVQ--
Rhopr-CRF/DH-R2 49 YGLCSELHTLTSNQILSDE--G-IIVQSTIYAVGYGFSLTALGLAVWIFLYY-------- L. stagnalis 44 ------PGSNESFVMPYDVPTGLICLLAFLYGSISLLAVIGNGLVILVIVKNRRMHTVTN D. melanogaster 17 -----GAEEEAEFERLYAAPAEIVALLSIFYGGISIVAVIGNTLVIWVVATTRQMRTVTN A. stephensi 21 -----DVRIVLEDENLYKVPIGLLVLLSIFYGTISILAVIGNSLVIWIVITTKQMQTITN A. aegypti 21 -----DVEIVKEQDALYDVPVGLVVLLSIFYGTISIIAVIGNSLVIWIVLTTKQMQTITN R. microplus 37 -----LPAANISSNKLYQVPVGFIVLLSIFYGIISLVAVAGNFMVMWIVATSRRMQTVTN RhoprKR 25 -----NQSVYFYEESLYEVPAGVIVLLSVFYGTISVVAVGGNFLVMWIVATSRRMQNVTN RhoprCAPA-R1 20 -----LEEYLIIVRGPKFLPLKILLPITFTYGILFISGLFGNLAVCIVIAYNKSMHNATN Rhopr-SKR-1 1 -------------MLPNESWWEAGKVQIPTYSIIFLLGLVGNILVILVLVKNKGMRTVTN Rhopr-SKR-2 11 -----PKSTSTTNTGGSDNGSGISELMIPLYMLIFILAVVGNSLVLATLTRNRRMRTVTN
Rhopr-CRF/DH-R2 98 ----------------------KDLWCLRNTIHTNLMCTYILAD-------LMWILSSIQ L. stagnalis 98 IFIPNLAVSDVIIGLFSIPFQFQAA-LLQRWVLANFMSSLPPFVQVVTVNLTIFTLRVIA D. melanogaster 72 MYIANLAFADVIIGLFCIPFQFQAA-LLQSWNLPWFMCSFCPFVQALSVNVSVFTLTAIA A. stephensi 76 MFIANLALADVTIGVFAIPFQFQAA-LLQRWNLPEFMCPFCPFVQLISVNVSVFTLTAIA A. aegypti 76 MFIANLALADVTIAVFAIPFQFQAA-VLQRWNLPEFMCPFCPFVQLLSVNVSVFTLTAIA R. microplus 92 FFIANLAVADIIIGLFSIPFQFQAA-LLQRWVLPEFMCAFCPFVQVLSVNVSIFTLTAIA RhoprKR 80 CFIANLALADIVIGLFAIPFQFQAA-LLQRWNLPNFMCPFCPFVQVLSVNVSVFTLTAIA RhoprCAPA-R1 75 YYLFSLAMSDLVLLLLGLPNDLSVFWQQYPWILGLLVCKLRALVSEMSSYVSVLTIVAFS Rhopr-SKR-1 48 VFLLNLAVSDILLGVLCMPFTLVGS-LLKDFVFGHFMCRLIPYMQACSVAVSGWTLVCLS Rhopr-SKR-2 66 VYLFNLAVADILLGVFCMPFTLIGQ-LLRNFVFGRIMCKLIPYFQAVSVSVAVWTLVAIS
Rhopr-CRF/DH-R2 129 VYVKTDPAICMVLFI-----LLHYLILTNYFWMFVEGLYL-------------------- L. stagnalis 157 --VDRYIAVIHPFKAGCSK--KRAAIIISIIWAVGIGAALPVPLFYWVEDLTE------- D. melanogaster 131 --IDRHRAIINPLRARPTK--FVSKFIIGGIWMLALLFAVPFAIAFRVEELTER-F-REN A. stephensi 135 --VDRHRAIINPLRARTSK--NISKFVISSIWMLSFVLAAPILFALRVRPVSYIALGGMN A. aegypti 135 --VDRHRAIINPLRARASK--NISKFVISAIWMMSFALAAPTLFALRVVPVSIVSLGETN R. microplus 151 --LDRYRAVMSPLKARTTK--LRAKFIICGIWTLAVAAALPCALALRVETQV-----E-- RhoprKR 139 --VDRHRAVLNPLSAPPSK--LRAKALLGAIWILAAILATPMAVALNVTYVE-----END RhoprCAPA-R1 135 --VERYTAICYPLKSYTTDKLNRVIKVIGTLWLISLGFAAPFAIYTTIDYVDFPPG---- Rhopr-SKR-1 107 --VERYYAICHPLRSRTWQTLTHAYRLIGAIWVCSLLLMTPISVLSELIPT---S----- Rhopr-SKR-2 125 --LERYFAICRPLKSRRWQTQFHAYKMIAIVWAMSLVWNSPILFVSRLLAM---GG----
Rhopr-CRF/DH-R2 164 ------------------------------------YMLVVETFTRENINLRAYLAIGWG L. stagnalis 206 ---NNIVIPRCDWHAPDNWLDFHLYYNTLLVCFQYLLPLVIITYCYCRIAWHIWGSRRPG D. melanogaster 185 NETYNVTRPFCMNKNL--SDDQLQSFRYTLVFVQYLVPFCVISFVYIQMAVRLWGTRAPG A. stephensi 191 DTYTNITVPFCKVVNF--EDGEILLYRYVLVLVQYFIPLFVISFVYIQMALRLWGSKTPG A. aegypti 191 ETYINMTKPFCQVVNF--EESEMLLYRYILTLVQYFVPLCVISFVYIQMALRLWGSKTPG R. microplus 200 SHALNLTKPFCHEVGI--SRKAWRIYNHVLVCLQYFFPLLTICFVYARMGLKLKESKSPG RhoprKR 190 HVGHVYTKPFCINTKL--SNNHMMAYRMILVSVQYLTPLCVISYAYAKMALRLWGSRAPG RhoprCAPA-R1 189 SGKAVIESAFCAMLKQ--NVPADVPLYELSCTLFFICPAVILIFLYVRIGL--------- Rhopr-SKR-1 157 GGHRK-----CRELWP--NEDIEKTYNLLLDFLLLVIPLIVMVTTYTLVAKTLWRVMKTQ Rhopr-SKR-2 176 KGRHK-----CREVWP--GRRSEGAYIIFLDIVLLMIPLLIMSLAYSLIVLKLWKGLQRE
Rhopr-CRF/DH-R2 188 IPVIIVIPSCLARA-------------------------------------FISDDYE-- L. stagnalis 263 ------A----------------------------------------------------- D. melanogaster 243 ------N----------------------------------------------------- A. stephensi 249 ------N----------------------------------------------------- A. aegypti 249 ------N----------------------------------------------------- R. microplus 258 ------N----------------------------------------------------- RhoprKR 248 ------N----------------------------------------------------- RhoprCAPA-R1 238 ------------------------------------------------------------ Rhopr-SKR-1 210 KPG---NEMGLK-------------------------------------------DNRVT Rhopr-SKR-2 229 LKH---SNSCLQTVDRSASLPTMTEVVISKNLNTNSEAIHRVIPADSQQGQFCNKNPVQM
Rhopr-CRF/DH-R2 209 YVL---------------------------ITKLRSSNNA-ETQQYRKATKALLVLIPLL L. stagnalis 264 ---------------------------------HVTTEDV-RGRNKRKVVKMMIIVVCLF D. melanogaster 244 ---------------------------------AQDSRDITLLKNKKKVIKMLIIVVIIF A. stephensi 250 ---------------------------------AQDSRDITMLKNKKKVIKMLIIVVALF A. aegypti 250 ---------------------------------AQDSRDMTMLKNKKKVIKMLIIVVALF R. microplus 259 ---------------------------------AQGARDAGILKNKKKVIKMLFVIVALF RhoprKR 249 ---------------------------------AQHSRDANLMRNKKKVIKMLVIVVALF
90
RhoprCAPA-R1 238 -----------TIKNNTKL-------------RGNVHGELQSIQSKKSIVSMLMAVVVAF Rhopr-SKR-1 224 WKQN---------SRG--------------SPHLRRSNTEKALKKKKRVVKMLFAVVLEF Rhopr-SKR-2 286 WLLKVKLEEGIAQRSGTPLEPLESPGPKFTRHAIRSNYMDKSIEAKKKVIRMLFVVVAEF
Rhopr-CRF/DH-R2 241 GVTYI-------LFIAGPTEGPYAYLFS---YIRAFLLSTQGLMVALLYCFLNTEVQNTV L. stagnalis 290 VLCWLPLQMYNLLHNINPLINHYHYINI-IWFSSNWLAMSNSCYNPFIYGLLNEKFKREF D. melanogaster 271 GLCWLPLQLYNILYVTIPEINDYHFISI-VWFCCDWLAMSNSCYNPFIYGIYNEKFKREF A. stephensi 277 GVCWFPLQLYNILHVTWPEINEYRFINI-IWFVCDWLAMSNSCYNPFIYGIYNEKFKREF A. aegypti 277 GICWFPLQLYNILHVTWSEVNEYRYINI-IWFVCDWLAMSNSCYNPFIYGIYNEKFKREF R. microplus 286 AFCWLPYQLYNILREVFPKIDKYKYINI-IWFCTHWLAMSNSCYNPFIYAIYNERFKREF RhoprKR 276 AICWLPLQTYNVLQDIFPQINGYRYINI-IWFCCDWLAMSNSCYNPFIYGIYNEKFKQEF RhoprCAPA-R1 274 FICWAPFHMQRLIYVYMSDYPWYGIVNVWLYYISGIFYYFSATINPILYNLMSLKYRKAF Rhopr-SKR-1 261 FVCWTPLYVINTITLFAPQAVYERLGYK-GISFLQLLAYSSSCCNPITYCFMNYRFRRAF Rhopr-SKR-2 346 FICWAPLHVLNTWYQFRPDLVHQYVGST-GVSLVQLLAYISSCCNPITYCFMNYRFRQAF
Rhopr-CRF/DH-R2 291 RHHFTRWKESRNLGARRYTCSRDWSPNTRTESVRLCSKHDVMPYRKRESVASENTTMTLV L. stagnalis 349 HQLFVMCPCWKARVDYYT-----EY---FSEDANICRRANTNGHCPANRHGAVGTTSTET D. melanogaster 330 NKRFAACFCKFKTSM-------------DAHERT------------FSMHTRASSIRSTY A. stephensi 336 RKRYPFKR-DQTYNHNHE------------SDKT------------SSIFTRVSSIRSTY A. aegypti 336 HKRYPFRGRNQSYHQEQL------------TDKT------------LSMFTRVSSIRSNY R. microplus 345 ATRCTCGGHRYKSPKS--------------------------------RFASYEQEDNST RhoprKR 335 QQRCPFSRRRKWTHGFGA-----GGSDSLDLDKT------------IHRFGSVNRNSSRW RhoprCAPA-R1 334 KQTLWCRKYNRIIKTPGL-----RETNSTSRQVNK-------SIKSMNMQHNQS----LL Rhopr-SKR-1 320 LKLFGCLREEKGSS---------------------------------------------- Rhopr-SKR-2 405 ISLFNFPRLCCWCGIPVE-----SKLAQRTDTAN----------EPNSLSANDS---TLY
Rhopr-CRF/DH-R2 351 GGSTNLARLS-------------------------------------------------- L. stagnalis 401 TRKSMLSR------------------------------------------SRCKGTRRR- D. melanogaster 365 ANSSMRIRSNLFGPARGGVNNGKPGLHMPRVHGSGANSGIYNGSSGQ--NNNVNGQHHQH A. stephensi 371 ATSSIRNKLST-NRYSASKQFKFPPPNHHFQHQPG---GHHNATGGAHLHELAFGTSK-- A. aegypti 372 ATSSIRNKLYT-GPIGGGSGN-------------G---GTHVGSGY---SSNAFYQNQNS R. microplus 373 IIVSM--------------RHS------FRLSFKN--SAPLKASTQV------------- RhoprKR 378 IRYSS--------------RVQYTPAQHYIYHCAN--SNTVHHSSQSEIEELCL------ RhoprCAPA-R1 378 ANN----------------------------------------------IED-------- Rhopr-SKR-1 ------------------------------------------------------------ Rhopr-SKR-2 447 AGRANRSEVMVLEK-----------------------------------EER--------
Rhopr-CRF/DH-R2 ------------------------------------------------------------ L. stagnalis 418 --RQTYDERRETSS---------------------------------------------- D. melanogaster 423 QSVVTFAATPGVSAPGVGVAMPPWR-----RNNFKPLHPNVIECEDDVAL------MELP A. stephensi 425 KGPVNFDGTVT---TTFATNHPREKKMDHRLVE---HDQLIASCIERLD-----HELACS A. aegypti 412 HHQQSYKSPNTNSVAGYQRNSTTDRNSSRKTAAGAPWDPKCCPCRQNSTRTSTAAASACP R. microplus ------------------------------------------------------------ RhoprKR ------------------------------------------------------------ RhoprCAPA-R1 384 ---IT------------------------------------------------------- Rhopr-SKR-1 ------------------------------------------------------------ Rhopr-SKR-2 464 ---V--------------------------------------------------------
Rhopr-CRF/DH-R2 ------------------------------------------------------------ L. stagnalis ------------------------------------------------------------ D. melanogaster 472 STTPPSEELASG-------------AGVQLALLSRESSSCICEQEFGSQTECDGTCILSE A. stephensi 474 STVDSSEDHRNGEPRTLNRPDIDGNGTGRAAKLRNGSS-----RE----CGLSIASNYAD A. aegypti 472 YRMPLPAVASDGDSGSEGGP-CNSAGGGQSPMINNDER-----QLLGADDNYGSAAQKLE R. microplus ------------------------------------------------------------ RhoprKR ------------------------------------------------------------ RhoprCAPA-R1 ------------------------------------------------------------ Rhopr-SKR-1 ------------------------------------------------------------ Rhopr-SKR-2 ------------------------------------------------------------
Rhopr-CRF/DH-R2 ----------------------------------------------------------- L. stagnalis ----------------------------------------------------------- D. melanogaster 519 VSRVHLPGSQAK----------------DKDAGKSLWQPL------------------- A. stephensi 525 RMALKHPHPDSGGESGDGEPKPGQRSSEERDSGGHLYCNDLEELGPYYD---------- A. aegypti 526 VISLDHPHPDSADDENGVAETPHSRTANGQEQDERLQLTSFISSGNGRHERFHFHINNL R. microplus ----------------------------------------------------------- RhoprKR ----------------------------------------------------------- RhoprCAPA-R1 ----------------------------------------------------------- Rhopr-SKR-1 ----------------------------------------------------------- Rhopr-SKR-2 -----------------------------------------------------------
91
Table 1: Similarity within transmembrane domains (TMs) of cloned invertebrate kinin receptors
based on sequence alignments carried out using the MUSCLE alignment tool and later scored on
the BLOSUM62 matrix.
TM1 TM2 TM3 TM4 TM5 TM6 TM7
Percent Pairwise Identity
71% 76% 75% 43% 52% 69% 87%
Percent Identical Sites
47% 56% 47% 21% 26% 50% 73%
92
Fig. 3: Expression profile of the RhoprKR transcript in the CNS and various feeding-related
tissues. Transcript expression is shown for the following tissues: CNS, fat body (FB), dorsal vessel
(DV), salivary glands (SG), foregut (FG), anterior midgut (AMG), posterior midgut (PMG),
hindgut (HG), and Malpighian tubules (MT). Transcript expression is relative to an average of the
three reference genes (α-tubulin, β-actin, ribosomal protein 49). Data represents the mean ± SEM
of three biological replicates per tissue, with three technical replicates for each biological replicate.
93
94
Fig. 4: The effects of RhoprKR transcript knockdown on R. prolixus hindgut contractions in
response to Rhopr-kinin 2 and the kinin analog 2139[F1]wp-2. Hindgut contractions were
performed on R. prolixus 7 days post-dsRNA injection. A) Changes in basal tonus in response to
varying concentrations of Rhopr-kinin 2 in dsARG-injected and dsRhoprKR-injected insects. B)
Changes in basal tonus in response to varying concentrations of 2139[F1]wp-2 in dsARG-injected
and dsRhoprKR-injected insects (One-way ANOVA followed by Tukey’s post-hoc test, *=p<0.05.
Data are means ± SEM of n = 5).
95
96
Fig. 5: The effects of RhoprKR transcript knockdown on in vivo feeding and diuresis in R. prolixus.
Weights of insects were measured after 20 minutes of feeding on saline supplemented with ATP
(time 0), and at 1 hour increments post-feeding for 4 hours. A) Weights of dsARG-injected and
dsRhoprKR-injected insects before and after artificial blood-feeding (time 0). B) The amount of
meal size times body weight consumed by dsARG-injected and dsRhoprKR-injected insects. C)
The rate of post-feeding diuresis of dsARG-injected and dsRhoprKR-injected insects. (Student’s
t-test, slopes tested for significance using an F-test. *=p<0.05. Data are means ± SEM of n=19-
23).
97
B)
A)
C)
98
Discussion
The R. prolixus kinin receptor (RhoprKR) has been successfully cloned, and its sequence,
structure, and expression profile analyzed. Examining the kinin signaling pathway within R.
prolixus is quite significant, due to its direct role in Chagas disease transmission (Bern et al., 2011).
For example, Rhopr-kinin 2, and its analog 2139[F1]wp-2 have been previously shown to be
potent in inducing hindgut contractions, a behaviour required for the excretion of urine (Bhatt et
al., 2014; see chapter 2).
Following analysis of its sequence and predicted structure, the RhoprKR transcript codes
for a kinin GPCR, belonging to the rhodopsin-like (family A) family of GPCRs. In comparison
with other invertebrate kinin receptors, there is a high degree of similarity between the sequences.
The predicted transmembrane domains within the kinin receptors are highly conserved in size and
spacing, but this conservation is not observed within TM4 and TM5, which was also observed by
Radford et al. (2004). The DRY/DRH/ERY motif is characteristic of all family A GPCRs which
is important in conformational changes for receptor activation (Capra et al., 2004; Rovati et al.,
2007). This motif is present as DRH within all of the insect kinin receptors analyzed, while in L.
stagnalis and R. microplus, it is present as DRY (Cox et al., 1997; Radford et al., 2002; Holmes et
al., 2002; Radford et al., 2004; Pietrantonio et al., 2005). In Rhopr-SKR-1, Rhopr-SKR-2 and
capa-r1 this motif exists as ERY (Paluzzi et al., 2010). The NPxxY domain within TM7, required
for inducing key structural changes within the receptor, is found in all family A GPCRs (Fritze et
al., 2003) and is observed in all of the kinin receptors as NPFIY but extends into the C-terminus
in R. prolixus and D. melanogaster. In addition, the first extracellular loop may be involved in the
binding of the kinin ligand, as it is also highly conserved (Radford et al., 2004). Disulphide bridges
are essential for the structural integrity of the receptor (Knudsen et al., 1997), with the Cys residues
99
that likely form this disulphide bridge conserved across all of the analyzed kinin receptors but is
not seen in the receptor sequence of the snail, L. stagnalis (Radford et al., 2004). Phosphorylation,
mediated by various protein kinases, are essential for various functions within GPCRs such as
internalization of the receptor and desensitization (Ferguson, 2001; Tobin, 2008). Ser and Thr
residues, which are common sites of phosphorylation by these kinases, appear to be fairly
conserved suggesting that these are common phosphorylation sites in kinin GPCRs (Radford et al.,
2004). In many invertebrate kinin receptors, receptor activation results in increases in intracellular
Ca2+ (O’ Donnell et al., 1998; Cady & Hagedorn, 1999; Tehrzaz et al., 1999; Cox et al., 1997). It
is expected that RhoprKR utilizes intracellular Ca2+ as a second messenger at the hindgut of R.
prolixus, but further functional studies need to be performed to confirm this.
The feeding strategy of R. prolixus requires the coordination of various neuroendocrine
factors to undergo post-prandial diuresis and excretion, and thereby rid the insect of excess water
and salts following a large blood meal (Orchard, 2006; Coast et al., 2002; Orchard, 2009). During
the production of primary urine, various diuretic and antidiuretic hormones act upon the anterior
midgut and MTs in order to facilitate the fluid and ion absorption and secretion required to
maintain osmotic balance (Coast et al., 2002; Maddrell, 1969). Kinins have been identified to have
myostimulatory effects on the anterior midgut but have no role in fluid transport within R. prolixus
(Te Brugge et al., 2009). An increase in the frequency of anterior midgut contraction that occurs
during feeding may assist in the mixing of the blood meal within the gut and flow of haemolymph,
thereby assisting in fluid absorption that occurs from the anterior midgut to the haemolymph
(Maddrell, 1964; Te Brugge et al., 2009). Expression of the RhoprKR transcript was observed in
the anterior midgut, which supports this myotropic role of Rhopr-kinins. While not directly
involved in the post-feeding diuresis or excretion, the salivary glands are responsible for the
100
secretion of substances during feeding into the host to ensure successful blood uptake, as these
substances prevent platelet aggregation, coagulation, and vasoconstriction within the host’s blood
vessels (see Orchard & Te Brugge, 2002). Kinins have been shown to induce dose-dependent
increases in salivary contractions, mediating the release of these substances during blood feeding
(Orchard & Te Brugge, 2002). As expected therefore, transcript expression of RhoprKR is seen
within the salivary glands.
Expression of the RhoprKR transcript was also observed in the foregut and posterior
midgut, and is higher in these tissues than that observed in the anterior midgut. During feeding,
the foregut functions to assist in the movement of the blood meal to the anterior midgut, where the
initial fluid absorption in the diuretic process occurs (Cooper & He, 1994; Te Brugge et al., 2009).
Like many other feeding and diuresis-related visceral muscle tissues, contraction and relaxation of
the foregut has been found to be under neuroendocrine control within various insects (Audsley &
Weaver, 2009). In Periplaneta americana, tachykinins were found to induce foregut contractions,
with proctolin and serotonin [5-hydroxytryptamine (5-HT)] inducing foregut contractions in
Teleogryllus commodus (Nässel et al., 1998; Cooper & He, 1994). In contrast, FGLamide-related
allostatins were shown to inhibit foregut contractions in L. maderae and L. migratoria (Duve et
al., 1995). The neuroactive chemicals that may be involved in modulating foregut contractions
have not been studied in R. prolixus. As expression of RhoprKR has been identified in the foregut,
Rhopr-kinins may also play a role in modulating movement of the foregut.
In contrast to the anterior midgut, the posterior midgut does not have a direct diuretic
function within R. prolixus. It instead aids in the digestion of the blood meal, as various digestive
enzymes have been identified within this tissue that aid in nutrient breakdown (Billingsley &
101
Downe, 1983). The transcript expression profile of RhoprKR suggests that Rhopr-kinins also act
upon the posterior midgut, possibly inducing contraction or aiding in blood digestion. As seen in
other tissues within R. prolixus, Rhopr-kinins likely elicit myotropic effects on this tissue. Kinin-
like immunoreactivity has been previously observed within endocrine cells in the posterior midgut
(Te Brugge et al., 2001). Within insects, contractions of the posterior midgut have also been found
to be under the influence of various neuroendocrine factors. Within Diploptera punctata, proctolin
and leucomyosuppressin alter contractions of the posterior midgut, albeit to a lesser degree than
the anterior midgut (Fusé & Orchard, 1998). Similarly, in Schiscostera migratoria, innervation
using FMRFamide-related peptides in seen in the posterior midgut, with SchistoFLRFamide,
leucomyosuppressin, and ManducaFLRFamide inhibiting spontaneous and proctolin-induced
contractions of midgut muscle (Lange & Orchard, 1998). Within D. melanogaster larvae,
immunoreactivity for a kinin receptor has also been observed in cells of the posterior midgut
(Veenstra, 2009), suggesting a possible role of kinins in posterior midgut function. The presence
of RhoprKR throughout the entire alimentary canal suggests that Rhopr-kinins may have a broader
role in feeding regulation. Relative to other analyzed tissues, expression levels of the RhoprKR
transcript was negligible in the MTs, fat body and dorsal vessel.
From an agrochemical perspective, identification of the RhoprKR sequence allows the
utilization of novel strategies to target the kinin signaling system within R. prolixus. Successful
RNAi-mediated knockdown of the RhoprKR transcript was found to have a significant impact on
hindgut contractions, as insects injected with dsRhoprKR had a reduced response to Rhopr-kinin
2 and the analog 2139[F1]wp-2. In addition, dsRhoprKR-injected insects had a significantly larger
meal compared to dsARG-injected insects, with a majority of dsRhoprKR-injected insects
consumed a meal that was over 10 times their body weight. These results suggest that Rhopr-kinins
102
may also function as satiety factors, as Aib-containing kinin analogs have been previously
described to induce antifeedant effects in R. prolixus (Lange et al., 2016). Within R. prolixus,
several neuroendocrine factors such as sulfakinin (Rhopr-SK-1) and corticotropin-releasing factor
(CRF)- like diuretic hormone (Rhopr-CRF/DH) have been identified in influencing feeding (Al-
Alkwai et al., 2017; Mollayeva et al., 2018) As co-localization of Rhopr-kinins and Rhopr-
CRF/DH has been observed within the CNS and midgut of R. prolixus, they may co-operatively
function to induce satiety following a blood meal (Te Brugge et al., 1999; Te Brugge et al., 2001;
Te Brugge et al., 2002; Mollayeva et al., 2018). However, despite the presence of kinin receptors
throughout the gut of R. prolixus, there were no significant differences in excretion rate between
dsRhoprKR-injected and dsARG-injected insects. The lack of significant differences on excretion
rate may be due to a partial knockdown of the RhoprKR transcript. The RhoprKR transcript is one
of many targets within R. prolixus that result in altered phenotypes due to the disruption of critical
signaling systems. Successful RNAi has been observed in a diverse set of tissues within R.
prolixus, all of which have resulted in physiological and behavioural changes that may interfere
with normal functioning. RNAi-mediated knockdown of the adipokinetic hormone receptor
resulted in an increased level of lipid content in the fat body of R. prolixus insects, and a decrease
in lipid levels in the haemolymph, due to the disruption of lipid mobilization (Zandawala et al.,
2015). The receptor for corazonin (CRZ), a neuropeptide implicated in dorsal vessel regulation in
insects, was targeted using RNAi, resulting in a reduced basal heartbeat, with a reduced response
to CRZ (Hamoudi et al., 2016). As the dorsal vessel is responsible for the circulation of
haemolymph within the insect, this knockdown can compromise the timely circulation of essential
endocrine factors. Development of novel pest-control strategies surrounding RNAi can prove to
be quite efficacious in preventing the spread of Chagas disease. On a larger scale, utilizing
symbiont-mediated RNAi can prove to be a successful next-generation pest control strategy
103
(Whitten et al., 2016). The efficacy of dsRhoprKR makes it an excellent candidate to be integrated
into essential gut bacteria, thereby knocking down signaling pathways that could ultimately
prevent the spread of the T. cruzi parasite through excretion (Whitten et al., 2016; Bern et al.,
2011).
In summary, based on its sequence similarity and expression profile, the RhoprKR
transcript encodes for an insect kinin receptor. Expression of RhoprKR in tissues such as the
foregut and posterior midgut indicate that Rhopr-kinins may have novel myotropic effects on these
tissues within R. prolixus. The effective knockdown of RhoprKR through RNAi, and the resultant
influence on kinin-stimulated hindgut contractions and feeding make dsRhoprKR a viable dsRNA
that can be incorporated in pest-control strategies, such as symbiont-mediated RNAi. Identification
of RhoprKR allows further investigation of the Rhopr-kinin signaling pathway, a neuropeptide
pathway that is closely associated with behaviours related to Chagas disease transmission.
104
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Supplementary Material
Supp. Fig. 1: Analysis of RhoprKR transcript expression following dsRhoprKR injection.
Reduction in transcript expression within the CNS and hindgut is shown for dsRhoprKR-injected
insects 2 days and 7 days post-dsRNA injection. Reduction in transcript expression is relative to
RhoprKR transcript expression in dsARG-injected insects. Data represents the mean ± SEM of
three biological replicates per tissue, with three technical replicates for each biological replicate.
Table S1: Primers used for the amplification of RhoprKR sequence fragments
Primer Name Primer Sequence (5’ to 3’)
KRFW1
GCGCAAATTGTTCATGGCTC
KRFW2
AAAGTCTGTACGAAGTTCCTGC
KRFW3
AAGCTTTATTGGGCGCTATTTG
KRFW4
GGTGGCTCTATTTGCAATATGT
KRFW5
AGCAAGAATTTCAACAACGATG
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KRRV1
GCAAAAAGGGCACATAAAATTT
KRRV2
CATTTAATGCTACAGCCATTGG
KRRV3
ATGCGTCTACTAGTGGCCAC
KRRV4
TGGCCGTCAGTGTGAATACA
KRRV5
GCCATTGGTGTTGCAAGGAT
Table S2: RACE Primers for the amplification of 5’ and 3’ regions of RhoprKR
Primer Name Primer Sequence (5’ to 3’)
KR5RACE1
ATTCTGCATGCGTCTACTAGTGGCCAC
KR5RACE2
TTACACCGGCAGGAACTTCGTACAGA
KR3RACE1
GTCCAGGTGCTCAGCGTCAATGTTAGT
KR3RACE2
GACGGCCATTGCTGTAGATAGACATCG
KR3RACE3
GCTATCCTTGCAACACCAATGGCTGTAG
Table S3: Primers used for RhoprKR dsRNA synthesis
Primer Name Primer Sequence (5’ to 3’)
dsRhoprKR
KRRNAIFW CGGCCTATTCGCTATACC
KRRNAIFWT7 TAATACGACTCACTATAGGGAGACGGCCTATTCGCTATACC
KRRNAIRV GGAATTACTCATTGCCAGCC
KRRNAIRVT7 TAATACGACTCACTATAGGGAGAGGAATTACTCATTGCCAGCC
dsARG
ARGFW AATAGTTTGCGCAACGTTG
112
ARGRV ATGAGTATTCAACATTTCCGTGTC
ARGFWT7 AATAGTTTGCGCAACGTTGTAATACGACTCACTATAGGGAGA
ARGRVT7 ATGAGTATTCAACATTTCCGTGTCTAATACGACTCACTATAGGGAGA
Table S4: Primers used for qPCR analysis
Primer Name Primer Sequence (5’ to 3’)
RhoprKR
KRQPCRFW TGCTCCACCCTCAAAATTAAGA
KRQPCRR ACCAACATGATCATTTTCTTCAACA
Rhopr-α-tubulin
FW GTGTTTGTTGATTTGGAACCTACAG
RV CCGTAATCAACAGACAATCTTTCC
Rhopr-β-actin
FW AGAGAAAAGATGACGCAGATAATGT
RV ATATCCCTAACAATTTCACGTTCG
Rhopr-ribosomal protein 49
FW GTGAAACTCAGGAGAAATTGGC
RV AGGACACATGCGTATC
113
Chapter 4 General Discussion
Feeding
The regulation of feeding behaviour is under the control of various neuropeptides within
insects (Audsley & Weaver, 2009), and here, the kinin and CAPA neuropeptides and their analogs
have been examined on feeding in Rhodnius prolixus
Kinin
Within R. prolixus, kinins are co-localized with corticotropin-releasing factor (CRF)- like
diuretic hormone (Rhopr-CRF/DH) within neurosecretory cells, with a decrease in staining at 2.5
hours after feeding (Te Brugge et al., 1999; Te Brugge et al., 2001; Te Brugge et al., 2002;
Mollayeva et al., 2018). As CRF/DH has been associated with feeding regulation in R. prolixus
(Mollayeva et al., 2018), these staining patterns suggested a role of kinins in feeding-related
behaviours. In Chapter 2, it was found that insects injected with the Aib-containing kinin analog
2139[Ф1]wp-2 took a significantly reduced blood meal relative to insects injected with a saline
control. These results complemented previous studies describing the antifeedant effects of Aib-
containing kinin analogs (Lange et al., 2016; Smagghe et al., 2010). This change in meal size may
be due to the disruption of feeding behaviour by interrupting the usual timely release of Rhopr-
kinins (Lange et al., 2016).
As transcript expression of RhoprKR was also confirmed in the hindgut (Chapter 3), these
receptors may be involved in modulating feeding behaviour, with the potent kinin analogs
interfering with this feeding signal (Te Brugge et al., 2001). In addition, dsRhoprKR-injected
insects consumed a significantly greater meal than dsARG-injected insects, suggesting that Rhopr-
114
kinins may possibly function as satiety factors (Chapter 3). Kinin-like immunoreactivity is present
in the abdominal nerves branching out to the abdominal regions, with additional peripheral neurons
that possess kinin-like staining within this area, suggesting the role of Rhopr-kinins in a gut-brain
feedback mechanism (Te Brugge et al., 2001). Neuropeptide release upon activation of a stretch
receptor has been previously demonstrated in crustaceans, as activation of a stretch-induced
mechanoreceptor in Homarus americanus resulted in a significant release of proctolin (Pasztor et
al., 1988). In D. melanogaster, the kinin pathway is implicated in meal regulation, as mutations in
the kinin/kinin receptor genes resulted in increased meal sizes and decreased meal frequency (Al-
Anzi et al., 2010). These phenotypic differences were due to impaired gut-brain signaling, as
neurons expressing the kinin and kinin receptor gene are expressed in the brain, with kinin-
containing neurons innervating the gut (Al-Anzi et al., 2010). A similar mechanism may exist in
R. prolixus, as kinin-like immunoreactivity and expression of the Rhopr-kinin transcript is present
within the CNS and digestive system (Te Brugge et al., 2001; Garima et al., 2014), with expression
of RhoprKR also seen within the CNS and digestive system (Chapter 3). More specifically, kinin-
like immunoreactivity has also been observed in neuropile throughout the CNS including the
subesophageal ganglion (SOG), which is primarily responsible for feeding regulation (Te Brugge
et al., 2001; Gllferin, 1972). Examining immunoreactivity of RhoprKR and transcript expression
through functional in situ hybridization (FISH) will assist in determining the specific localization
of RhoprKR within the CNS and digestive tissues. This will determine whether RhoprKR also
exists within these peripheral neurons near the abdomen, and whether similar staining patterns are
observed that have been seen with kinins.
115
CAPA
To further examine the neuropeptides that may modulate feeding behaviour, RhoprCAPA-
2 and its analog were also examined on in vivo feeding behaviour. As RhoprCAPA-2 is responsible
for the cessation of diuresis (Paluzzi et al., 2008; Paluzzi et al., 2010), it’s role in feeding regulation
was examined. R. prolixus that were injected with the CAPA analog 2129-SP3[Ф3]wp-2 had a
significantly higher blood-meal relative to the saline control (Chapter 2). As the CAPA analog is
an antagonist to the CAPA receptor (Jiang et al., 2015; Alford et al., 2019), this increase in blood
meal size suggest that RhoprCAPA-2 may have an additional role in serving as a feeding signal.
RhoprCAPA-2 appears to be released three to four hours after feeding, coinciding with the end of
diuresis (Orchard & Paluzzi, 2009). CAPA may activate a signaling cascade that possibly induces
satiety, or prevents additional feeding events. In addition, this feeding signal can be examined
through injection of the CAPA analog a few days following a blood meal, or at different states of
satiety and assessing feeding behaviour, as R. prolixus insects only feed once during each instar
(Buxton, 1930).
Following in vivo feeding assays, in vitro physiological assays on relevant feeding and
diuresis-related tissues were performed with the kinin and CAPA neuropeptides and their analogs
to uncover their role in these behaviours, and the efficacy of the analogs on the tissues.
Myotropic effects
Kinin
Rhopr-kinins, and Aib-containing kinin analogs have been previously shown to stimulate
hindgut contractions in R. prolixus (Te Brugge et al., 2002; Bhatt et al., 2014). These kinin analogs
are more biologically active in inducing hindgut contractions than the endogenous Rhopr-kinins
116
(Bhatt et al., 2014). In Chapter 2, the Aib-containing analog 2139[Ф1]wp-2 was potent in inducing
hindgut contractions. Following RNAi-mediated knockdown of the RhoprKR transcript, the
effects of Rhopr-kinin 2 and kinin analog were also examined (Chapter 3). The hindguts from
dsRhoprKR-injected insects had a reduced response to Rhopr-kinin 2 and 2139[Ф1]wp-2, but
2139[Ф1]wp-2 was still found to induce stronger hindgut contractions than Rhopr-kinin 2.
Analysis of the expression profile of the RhoprKR transcript showed high expression levels in the
hindgut, supporting the likelihood that RhoprKR indeed codes for a kinin receptor (Chapter 3).
In addition to its myotropic effects on the hindgut, kinins also stimulate contractions of the
anterior midgut and salivary glands, highlighting the multifunctional role of kinins in feeding and
diuresis-related tissues (Orchard & Te Brugge, 2002; Te Brugge et al., 2009). As the Aib-
containing kinin was potent in inducing hindgut contractions (Chapter 2, 3) examining its effects
on the anterior midgut and salivary glands would assist in determining their efficacy to be used as
lead compounds.
Following analysis of the expression profile of RhoprKR, high expression was also
confirmed in the foregut and posterior midgut (Chapter 3). However, the physiological role of
Rhopr-kinins on the foregut and posterior midgut have not been thoroughly examined. Within
insects, contractions of the foregut and posterior midgut are regulated by a host of neuropeptides
(Audsley & Weaver, 2009). The presence of RhoprKR within these tissues suggests that Rhopr-
kinins may regulate the alimentary canal as a whole, in turn regulating various aspects of the
digestive process following blood-feeding (Chapter 3). The kinin-like staining patterns support the
likelihood that Rhopr-kinins modulate foregut and posterior midgut function, as kinin-like
immunoreactivity has been detected in endocrine cells within the posterior midgut, and corpus
117
cardiacum (CC) (Te Brugge et al., 2001). The CC serves as a neurohemal organ that is located
above the foregut, therefore Rhopr-kinins may be released into the haemolymph as neurohormones
to modulate foregut contractions and salivary gland muscle contractions. (Orchard & Te Brugge,
2002).
CAPA
CAPA neuropeptides have been studied extensively across insects and elicit both
myotropic and diuretic effects (Predel & Wegener, 2006). As transcript expression of a
RhoprCAPA receptor was detected in the hindgut (Paluzzi et al., 2010), the effects of the
RhoprCAPA neuropeptides (RhoprCAPA-1, RhoprCAPA-2, Rhopr-pk1) were examined on
hindgut contractions (Chapter 2). Interestingly, in the insect species studied, CAPAs have shown
to be mutually exclusive in their myotropic or diuretic effects (Predel & Wegener, 2006). In R.
prolixus, none of the three CAPA neuropeptides on their own yielded any changes in hindgut
contractions (Chapter 2).
Within R. prolixus, various neuroendocrine factors have been found to act in concert to
elicit changes in basal tonus of the hindgut (Bhatt et al., 2014; Haddad et al., 2018). This was seen
with RhopCAPA-2 and Rhopr-kinin 2, as RhoprCAPA-2 potentiated the excitatory effects of
Rhoprkinin-2 (Chapter 2). It is possible that this interaction may occur at the intracellular level,
through separate second messenger pathways. Activation of the CAPA receptor results in increases
in cGMP, which in turn activates a phosphodiesterase that degrades cGMP (Paluzzi & Orchard,
2006; Paluzzi et al., 2013). In invertebrates, kinin receptors have been shown to cause intracellular
Ca2+ increases through the inositol triphosphate (IP3) pathway (O’ Donnell et al., 1998; Cady &
Hagedorn, 1999; Tehrzaz et al., 1999; Cox et al., 1997). It is likely that RhoprKR activates this
118
second messenger pathway also, due to the high sequence similarity in comparison to other
invertebrate kinin receptors (Chapter 3). It is also hypothesized that the CAPA receptor may be
constitutively active, stably increasing levels of cGMP and decreasing levels of cAMP (Chapter
2), with the CAPA analog serving as an inverse agonist. Serotonin [5-hydroxytryptamine (5-HT)],
which utilizes cAMP as a second messenger, causes increases in the frequency of hindgut
contractions (Orchard, 2006). The CAPA analog was found to potentiate the effects of 5-HT on
contraction frequency, while RhoprCAPA-2 inhibited them. The inhibitory effects of
RhoprCAPA-2 may be due to the elevated levels of cGMP, in turn activating the phosphodiesterase
that degrades cAMP, thereby reducing muscle stimulation. Figures 1,2, and 3 are models proposing
the mechanism of these intracellular interactions. Within invertebrates, constitutively active
GPCRs modulate a host of behaviours. In Caenorhabditis elegans, a constitutively active 5-HT7-
like GPCR is involved in pharyngeal pumping, while in Schistosoma mansoni, a constitutively
active acetycholine GPCR regulates larval motility (Hobson et al., 2003; MacDonald et al., 2015).
In addition, interaction of separate second messenger pathways has been found to occur in insects,
as cAMP potentiates the release of intracellular Ca2+ from the endoplasmic reticulum through the
IP3 pathway in the salivary glands of Calliphora vicina (Schmidt et al., 2008).
To further investigate this interaction, a calcium mobilization assay must be performed to
confirm RhoprKR ligand binding and its associated second messenger pathway. Utilizing
HEK293/CNG cells will assist in determining whether luminescence that occurs through activation
of RhoprKR is through the activation of cytosolic cAMP or intracellular Ca2+ through the IP3
pathway. Measuring levels of intracellular Ca2+ through in vivo calcium imaging (Russell, 2011)
during sole application of Rhopr-kinin 2 and a mixture of Rhopr-kinin 2 and RhoprCAPA-2 will
assist in determining whether higher levels of intracellular Ca2+ are achieved during this
119
interaction. In addition, measuring levels of cGMP via cGMP radioimmunoassay (Paluzzi &
Orchard, 2006) within the hindgut upon application of the CAPA analog will confirm whether this
analog is functioning as an inverse agonist on CAPA receptors within the hindgut.
120
Fig. 1: Model describing the possible second messenger pathways associated with the activation
of the R. prolixus kinin and 5-HT receptors, and the constitutively active CAPA receptor in the
hindgut
A) Activation of the kinin receptor increases hindgut contractions through intracellular Ca2+
increases mediated by the IP3 pathway (PIP2: Phosphatidylinositol 4,5-bisphosphate; PLC:
phospholipase C)
B) Activation of the 5-HT receptor stimulates increases in frequency of hindgut contraction
through activation of an adenyl cyclase (AC) causing increases in cAMP, resulting in activation
of protein kinase A (PKA) and eliciting further downstream effects leading to hindgut contraction
stimulation.
C) A constitutively active CAPA receptor stably increases cGMP levels, activating a
phosphodiesterase (PDE) that degrades cAMP D) Activation of the CAPA receptor through
RhoprCAPA-2 (agonist) contributes to greater increases in cGMP content, resulting in more cAMP
degradation.
121
√β
U U U
U U U
Y
Ca2+
[cGMP]
Endoplasmic Reticulum
α
CAPA Receptor(Constitutively
active)
[cAMP]PDE
5-HT (Agonist)
√
Ca2+
Endoplasmic Reticulum
Increase in Frequency of Hindgut Contraction
[cAMP] ATP
AC
αGTP
β
PKA
√
U U U
U U U
Y
√β
U U U
U U U
Y
Ca2+
[cGMP]
Endoplasmic Reticulum
α
[cAMP]PDE
RhoprCAPA-2 (Agonist)
Ca2+
Endoplasmic Reticulum
√β
Y U U U
U U U PIP2
IP3IP3 SensitiveCa2+ Channel
αGTP
Rhopr-kinin 2 (Agonist)
Increase in Hindgut Contraction
PLC
B)
C)
D)
A)
122
Fig. 2: Model describing the possible intracellular interactions between RhoprCAPA-2 and Rhopr-
kinin 2 and 5-HT in the hindgut of R. prolixus
A) Activation of the CAPA receptor by RhoprCAPA-2 causes increases in cGMP levels, which
interact with the IP3 pathway (activated by Rhopr-kinin 2), causing potentiated increases in hindgut
contraction.
B) Activation of the CAPA receptor by RhoprCAPA-2 causes increases in cGMP levels, thereby
decreasing cAMP levels through action of a phosphodiesterase, in turn reducing the stimulatory
effects of 5-HT on the frequency of hindgut contractions.
123
IP3 SensitiveCa2+ Channel
[cGMP]
Ca2+
Endoplasmic Reticulum
√β
Y U U U
U U U PIP2
IP3
αGTP
Rhopr-kinin 2 (Agonist)
PLC
Potentiated Increase in Hindgut Contraction
√β
U U U
U U U
Y
α
RhoprCAPA-2 (Agonist)
5-HT (Agonist)
√
Ca2+
Endoplasmic Reticulum
Reduction of 5-HT-Stimulated Increase in Frequency of
Hindgut Contraction
[cAMP] ATP
AC
αGTP
β
PKA
√
U U U
U U U
Y
[cGMP]
√β
U U U
U U U
Y
α
RhoprCAPA-2 (Agonist)
PDE
A)
B)
124
Fig. 3: Model describing the possible effects of the CAPA analog 2129-SP3[Ф3]wp-2 and its
intracellular interactions with Rhopr-kinin 2 and 5-HT in the hindgut of R. prolixus
A) The CAPA analog acts as an inverse agonist on the CAPA receptor, resulting in decreased
levels of cGMP, thereby preventing the degradation of cAMP and resulting in increases of hindgut
contraction.
B) The CAPA analog decreases cGMP levels, preventing an interaction with the IP3 pathway
(activated by Rhopr-kinin 2), resulting in no potentiation
C) The CAPA analog decreases cGMP levels, thereby preventing degradation of cAMP by a
phosphodiesterase, increasing cAMP levels, and potentiating the 5-HT-stimulated increases in the
frequency of hindgut contractions.
125
[cGMP]
√β
U U U
U U U
Y
Ca2+
Endoplasmic Reticulum
α
2129-SP3[F3]wp-2 (Inverse agonist)
[cAMP]PDE
PKA
Increase in Hindgut Contraction
[cGMP]
IP3 SensitiveCa2+ Channel
Ca2+
Endoplasmic Reticulum
√β
Y U U U
U U U PIP2
IP3
αGTP
Rhopr-kinin 2 (Agonist)
PLC
No Potentiated Increase in Hindgut Contraction
√β
U U U
U U U
Y
α
2129-SP3[F3]wp-2 (Inverse agonist)
5-HT (Agonist)
√
Ca2+
Endoplasmic Reticulum
Potentiated Increases in Frequency of Hindgut
Contraction
[cAMP] ATP
AC
αGTP
β
PKA
√
U U U
U U U
Y
[cGMP]
√β
U U U
U U U
Y
α
2129-SP3[F3]wp-2 (Inverse agonist)
PDE
A)
B)
C)
126
Diuretic effects
As RhoprCAPA-2 was identified as the first anti-diuretic hormone in R. prolixus, the
effects of the CAPA analog were tested on MT fluid secretion (Paluzzi & Orchard, 2006; Paluzzi
et al., 2008). Interestingly, Rhopr-kinins have no role in diuresis within R. prolixus, in contrast to
their diuretic effects in many other insects (Torfs et al., 1999). The CAPA analog was found to
prevent the inhibitory effects of RhoprCAPA-2 on 5-HT-stimulated tubules, supporting its
function as a CAPA receptor antagonist within the MTs (Chapter 2). The in vitro effects of the
CAPA analog support the differences in the rate of diuresis that was observed in R. prolixus insects
injected with the CAPA analog, as they had a significantly greater rate of post-feeding diuresis in
the first hour (Chapter 2). These effects of the CAPA analog highlight its promise as a lead
compound for pest-control development, as it can induce excess fluid loss in R. prolixus by
preventing the anti-diuretic signal of RhoprCAPA-2. As RhoprCAPA-2 also abolishes the
synergism that occurs between 5-HT and Rhopr-CRF/DH on stimulating upper MT secretion
(Paluzzi et al., 2011), examining the CAPA analog’s effects on the abolishment of this synergism
would provide more insight on its efficacy. In addition to the MTs, RhoprCAPA-2 has anti-diuretic
effects on the anterior midgut, as it inhibits 5-HT-stimulated fluid absorption (Ianoswki et al.,
2010). This would be an additional tissue that the CAPA analog can be tested on for its effects as
an antagonist.
Figure 4 provides a summary of the roles of kinin and CAPA in R. prolixus feeding and
diuresis-related behaviours.
127
Fig 4: The physiological roles of kinin and CAPA in the alimentary canal of R. prolixus.
Highlighted regions represent novel research findings.
128
ForegutRhoprKR expression confirmedUnknown physiological role
Salivary GlandsRhoprKR expression confirmedIncreases basal tonus (Orchard & Te Brugge, 2002)
Anterior MidgutRhoprKR expression confirmedIncreases frequency of contractions (Te Brugge et al., 2009)
Posterior MidgutRhoprKR expression confirmedUnknown physiological role
HindgutRhoprKR expression confirmedIncreases basal tonus, frequency of contractions (Te Brugge et al., 2002; Garima et al., 2014)
Anterior Midgutcapa-r expression confirmed (Paluzzi et al., 2010)Downregulates 5-HT-stimulated fluid absorption (Ianowski et al., 2010)
KininCAPA
Upper Malpighian Tubulescapa-r expression confirmed (Paluzzi et al., 2010) Inhibits 5-HT-stimulated fluid secretion (Paluzzi et al., 2008)Abolishes 5-HT/DH synergism (Paluzzi et al., 2011)
Hindgutcapa-r expression confirmed (Paluzzi et al., 2010) RhoprCAPA-2 potentiates Rhopr-kinin 2 hindgut contraction stimulation(no individual effects of RhoprCAPA neuropeptides)
Lower Malpighian Tubulescapa-r expression confirmed (Paluzzi et al., 2010)Unknown physiological role
129
Summary of Physiological Effects of Analogs
Following the investigation of the neuropeptide analogs on feeding and diuresis-related
behaviours, they were found to cause potent changes in physiology that can potentially interfere
with the normal functioning of R. prolixus. The kinin analog 2139[Ф1]wp-2 stimulated dose-
dependent increases in basal tonus of the hindgut and was more effective than the endogenous
Rhopr-kinin 2, and also had anti-feedant effects. The CAPA analog 2129-SP3[Ф3]wp-2 also
stimulated dose-dependent increases in hindgut contraction and potentiated the effects of 5-HT on
the frequency of hindgut contractions (not seen by endogenous RhoprCAPAs), increases the size
of blood meals, and prevented RhoprCAPA-2’s inhibition on 5-HT stimulated MTs.
The objective of this research project was to investigate the roles of Rhopr-kinins and
RhoprCAPAs on the feeding and post-feeding physiology of R. prolixus. Analogs that were
developed for kinin and CAPA were assessed for their efficacy to be used as compounds in pest-
control development. The neuropeptide analogs had a significant impact in vivo and in vitro and
show promise for use in various pest-control strategies. Novel roles of CAPA and kinins within
feeding, diuresis and excretion have been uncovered using a physiological and molecular
approach. Lastly, RhoprKR has been successfully identified, cloned and sequenced, showing high
susceptibility to RNAi-mediated knockdown. Table 1 provides a summary of the future directions
in light of these research findings. Overall, this thesis highlights the importance of neuropeptide
signaling within R. prolixus, and its role in physiology and behaviours related to disease
transmission.
130
Table 1: A summary of future directions
Physiological Approach Molecular Approach Investigate the effects on 2139[Ф1]wp-2 on contractions of salivary glands and anterior
midgut
Functionally characterize RhoprKR and confirm its second messenger pathway
Investigate the effects of 2129-SP3[Ф3]wp-2 on anterior midgut absorption, and 5-HT/DH
synergism in upper MTs
Perform FISH to determine RhoprKR transcript localization
Measure levels of second messengers involved in Rhopr-kinin/RhoprCAPA
interaction within the hindgut
Using functional receptor assay to determine the binding efficiency of the kinin analogs and any possible interference by the CAPA
neuropeptides and the CAPA analog Perform immunohistochemistry for
RhoprKR-like staining in the CNS and digestive system
Perform additional immunohistochemistry for RhoprCAPA-2 like innervation
Examine effects of RhoprCAPA-2 on various states of satiation during R. prolixus life cycle
131
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