Molecular Characterization, Expression Analysis, and Physiological Roles of

Allatotropin in Rhodnius prolixus

by

Maryam Masood

A thesis submitted in conformity with the requirements

for the degree of Master of Science

Department of Cell and Systems Biology

University of Toronto

© Copyright by Maryam Masood 2013

ii

Molecular Characterization, Expression Analysis, and Physiological Roles of Allatotropin

in Rhodnius prolixus

Maryam Masood

Master of Science

Department of Cell and Systems Biology

University of Toronto

2013

ABSTRACT

Rhodnius prolixus, the principal Chagas disease vector, requires a blood meal to complete its

moult cycle into the next stage. Allatotropins (ATs), a family of peptides first isolated from

Manduca sexta, have been shown to regulate the biosynthesis of juvenile hormone, an insect

growth and development hormone; however, ATs, being multimodal peptides, also exhibit

myotropic effects on some insect visceral muscles. Here, this AT family of peptides has been

examined in R. prolixus. Genomic analysis revealed a cDNA fragment of 973bp encoding one

mature amidated AT tridecapeptide (Rhopr-AT) with high transcript levels observed, via RT-

PCR, in the central nervous system (CNS) and pool of fat body and trachea. AT-like

immunoreactive neurons were found throughout the CNS and AT-like immunoreactive

processes were present on some peripheral tissues. Bioassays using hindgut and dorsal vessel

contraction, however, failed to demonstrate any effect of Rhopr-AT on these tissues. Future

work will examine the effects of Rhopr-AT on JH production.

iii

Acknowledgments

I would like to express a heartfelt gratitude to my supervisor, Dr. Orchard, whose expertise,

understanding, patience, and vast knowledge added considerably to my graduate experience. I

appreciate his ongoing guidance and advice that enabled me to stay on track and work at a

smooth pace throughout the Master’s program. I am also extremely thankful to Dr. Angela

Lange and Dr. Joel Levine for providing me with invaluable input and feedback that helped

shape me into a better research student. Furthermore, I am indebted to my colleagues without

whom this journey would be incomplete. Meet, Dohee, Lisa, Nikki, Laura, Himali all the

undergraduates and work-study students, thank you all for enriching my graduate experience!

I feel special thanks is in order for Garima and Nigam for their continual guidance, support and

mere presence with which they brought the lab to life and made it such a wonderful place to be

in! Thank you amigos!

On a final note, I’d like to thank my family for putting up with my “princess” behaviour over

the last two years. Your support was absolutely fundamental to my research and I am extremely

appreciative!

iv

Table of Contents ABSTRACT ................................................................................................................................... ii

Acknowledgments ......................................................................................................................... iii

List of Tables ................................................................................................................................... v

List of Figures ................................................................................................................................. vi

List of In-text Abbreviations ........................................................................................................ vii

Molecular Characterization, Expression Analysis and Physiological Roles of Allatotropin

in Rhodnius prolixus ...................................................................................................................... 1

INTRODUCTION ........................................................................................................................... 1

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

Feeding-related physiological processes in Rhodnius prolixus .................................................. 2

Neuropeptides and Families ........................................................................................................ 3

G-protein coupled receptors (GPCRs) ........................................................................................ 5

Allatoregulatory Peptides ............................................................................................................ 6

Allatostatins ................................................................................................................................. 7

Allatotropins ................................................................................................................................ 9

THESIS OBJECTIVE ................................................................................................................... 16

MATERIALS AND METHODS .................................................................................................. 17

RESULTS ...................................................................................................................................... 27

DISCUSSION ................................................................................................................................ 54

FUTURE DIRECTIONS ............................................................................................................... 62

REFERENCES .............................................................................................................................. 64

v

List of Tables

Table 1. Sequences of known or predicted allatotropin peptides.. ................................................ 10

Table 2. Sequence similarities between allatotropin-like peptides of different lepidopteran

species ............................................................................................................................................ 14

Table 3. Gene-specific primers used for Rhopr-AT prepropeptide cDNA amplification ............. 19

Table 4. Amino acid sequence and purity of peptides used for the bioassays............................... 25

vi

List of Figures

Figure 1. A schematic representing the possible targets of allatoregulatory peptides..................... 8

Figure 2. A) Nucleotide sequence of allatotropin (AT) precursor peptide cDNA in

R. prolixus. B) Schematic representation of Rhopr-AT gene organization. C) Northern

blot of Rhopr-AT transcript ........................................................................................................... 28

Figure 3. ClustalO alignment of Rhopr-AT prepropeptide sequence with allatotropin

prepropeptide sequences from other arthropods. ........................................................................... 31

Figure 4. Phylogenetic relationship of allatotropin prepropeptide sequence with

other insect sequences.................................................................................................................... 33

Figure 5. RT-PCR representing the expression profile of the Rhopr-AT transcript in

R. prolixus ...................................................................................................................................... 35

Figure 6. Confocal images showing AT-like immunostaining in cells of R. prolixus

CNS, corpus cardiacum and dorsal vessel ..................................................................................... 39

Figure 7. Confocal images showing AT-like immunostaining in cells of R. prolixus

CNS and peripheral nerves ............................................................................................................ 41

Figure 8. Camera lucida map depicting distribution of AT-like immunoreactive cells

within the CNS. A) Dorsal view. B) Ventral view of the brain..................................................... 43

Figure 9. Confocal images showing AT-like immunoreactivity in peripheral tissues

of R. prolixus ................................................................................................................................. 45

Figure 10. Sample traces depicting effects of Rhopr-AT and Rhopr- Kinin 2 on IVth

(A) and Vth

(B) instar hindguts ...................................................................................................... 48

Figure 11. Dose-response curves representing the change in hindgut contractions of

IVth

instar R. prolixus hindguts ...................................................................................................... 50

Figure 12. A) Dose-response curve for Rhopr-AT activity on heart-beat of R. prolixus .............. 52

vii

List of In-text Abbreviations

Note: Only abbreviated scientific terms used repeatedly have been listed here. Initial use of an

in-text abbreviation is preceded by the full form.

AST: Allatostatin

AT: Allatotropin

AT-like-ir: allatotropin-like immunoreactive

cAMP: 3’-5’-cyclic adenosine monophosphate

CC/ CA: corpora cardiaca/ corpora allata

CNS: central nervous system

CRF: corticotropin releasing factor

DH: diuretic hormone

DV: Dorsal vessel

GPCR: G protein-coupled receptor

HG: Hindgut

JH: Juvenile Hormone

MALDI-TOF MS/MS: matrix assisted laser desorption ionization time-of-flight tandem

mass spectrometry

MTGM: mesothoracic ganglionic mass

MTs: Malpighian tubules

ORF: open reading frame

PRO: prothoracic ganglion

SG: Salivary gland

SOG: suboesophageal ganglion

SP: Sex peptide

Rhopr-kinin 2: Rhodnius prolixus kinin- 2

UTR: Untranslated region

1

Molecular Characterization, Expression Analysis and Physiological Roles of

Allatotropin in Rhodnius prolixus

INTRODUCTION

Rhodnius prolixus

American trypanosomiasis also referred to as the Chagas disease is a widespread pandemic in

Central and South America. Approximately 7-8 million people are infected worldwide, though

mostly in Latin America, with an estimated report of 50,000 deaths each year (Lima et al., 2010;

Rassi et al., 2010; World Health Organization, 2012). Rhodnius prolixus, one of the principal

vectors for Trypanosoma cruzi - the parasite responsible for Chagas disease, is an obligate

feeder whereby each blood meal initiates growth and development into the next instar stage.

Since the lifecycle of R. prolixus is dependent on the availability of a blood meal, each instar,

when provided the opportunity, is capable of gorging up to ten times its original body weight.

The considerable intake of fluid limits mobility, and increases the risk of predation; therefore,

within 2-3 minutes of blood feeding R. prolixus urinates and it is during this post-prandial

diuresis that the parasite, T. cruzi, is transmitted from the vector to the host (Orchard, 2006;

Rassi et al., 2010). Transmitted T. cruzi leads to either an acute symptomatic phase, expressed

by 1-2 % of the population, or a chronic asymptomatic phase, expressed by about 70% of the

infected population (World Health Organization, 2012). The acute phase is treatable with

antiparastic drugs such as benznidazole (Rassi et al., 2010); however, since the symptoms are

mild they usually go unnoticed and the disease develops into a fatal phase leading to chronic

cardiomyopathy, gastrointestinal and neurological problems (Rassi et al., 2010; World Health

Organization, 2012). Since the chronic stage of Chagas disease is incurable, the only other

treatment currently available is prevention. Triatomes such as R. prolixus are commonly found

in thatched roofs along with nooks and crevices associated with poor housing conditions (World

Health Organization, 2008). Therefore, Chagas disease has been seen to be strongly associated

with socio-economic conditions, with people at the poverty line at higher risk of infection.

Control programs implemented in regions of Central and South America have been successful in

diminishing the number of disease cases since the late 1990s (Rassi et al., 2010); however,

2

complete eradication can only be possible by inhibition of the vector host and information

pertinent to their lifecycles can be aided by relevant molecular and physiological research.

Feeding related physiological processes in Rhodnius prolixus

Diuresis

Rhodnius prolixus is capable of engorging a blood meal up to 10-12 times its original unfed

body weight. The substantial blood meal causes inflation of the abdomen leading to activation

of stretch receptors that trigger a cascade of events regulated by neurohormones and hormones;

diuresis being one of them. The imbibed meal is rich in water and ions which are consequently

absorbed from the gut into the haemolymph and hence need to be sequentially secreted from the

insect to maintain the internal osmotic balance of the haemolymph (Coast, 2009). The amount of

water secreted in the urine depends on the pre-feeding state of the insect, with dessicated insects

secreting less volume of urine compared to newly emerged fed instars (Maddrell, 1964). Ionic

regulation is controlled by specialised Malpighian tubules (MTs) that are characterised by their

location and relative epithelial conductance to ions; the upper tubules secrete excess Na+ ions

while the lower tubules preferentially reabsorb K+ and Cl

- ions (Coast, 2009). Thus, eventually,

a hyposomotic urine is produced which travels to the hindgut to be released from the anus.

Growth and development

Another physiological response initiated by abdominal stretch due to feeding is initiation and

completion of the moult cycle (Callier and Nijhout, 2013). Wigglesworth’s pioneering studies

(see 1934, 1948) regarding ecdysis identified the presence of a moulting hormone, later referred

to as “juvenile hormone” (JH) that was released from the corpora allata (CA) of R. prolixus.

Regulated production and release of this moulting hormone during critical time periods was

shown to result in either ecdysis into the next instar or metamorphosis into adults. However,

since then it has been determined that ecdysis is not an isolated event controlled by a single

hormone, rather a cocktail of hormones orchestrate growth and development and ecdysis

behaviour. Some of the neurohormones and hormones known to be involved in other insects

regulating this process are ecdysteroids, corazonin, preecdysis-triggering hormone (PETH),

ecdysis triggering hormone (ETH), eclosion hormone (EH) and crustacean cardioactive peptide

(CCAP) (Kim et al., 2006a; Lee do and Lange, 2011; Roller et al., 2010; Vafopoulou et al.,

3

2012; Zitnan and Adams, 2005). Ecdysis related hormones are highly conserved among insect

species and orthologues are seen in Diptera and Lepidopterans (Kim et al., 2004).

Neuropeptides and Families

Physiological processes including foraging, feeding, growth, development, metabolite

regulation, digestion, reproduction and cardiovascular control etc are modulated by numerous

neuropeptides that are released in a developmental stage-specific manner (for specific roles of

neuropeptides on aforementioned physiological processes, see: Audsley and Weaver, 2009;

Coast, 1996, 2009; Mykles et al., 2010; Wielendaele et al., 2013). Neuropeptides are short

chains of amino acids that offer great flexibility in function through structure and mode of

action; they may be neuromodulators, neurohormones or neurotransmitters (Orchard et al.,

2001). Neurohormones are neuropeptides (or amines) released into the haemolymph, from

neurosecretory cells, that subsequently circulate throughout the body; hence targeting multiple

tissues simultaneously. A neurohormone-mediated response is elicited only by those tissues that

possess appropriate ligand-specific receptors. Neurotransmitters, on the other hand, are released

at synapses, and result in discreet, private communication; while neuromodulators are released

locally in a paracrine manner and diffuse locally towards target tissues (Nässel, 2002; Orchard

et al., 2001). Due to the relative modes of transfer the duration of the message also differs

between neurotransmitters and neurohormones, with neurohormones relaying persistent, long-

term messages and neurotransmitters conveying short-term messages (Nässel, 2002; Orchard et

al., 2001). The multiplicity of neuropeptides and their diverse physiological roles make them

excellent cell-cell communication molecules; hence neuropeptides are well-represented among

various taxa, with even simple organisms such as coelenterates and nematodes producing

several neuropeptides that work in concert with one another.

Neuropeptides act through multiple receptor types, though most interact with G-protein coupled

receptors (GPCRs), and functional flexibility is further enhanced by tissue receptivity, i.e.

receptors for neuropeptides are expressed in varying amounts in different tissues in a stage-

specific manner. In Aedes aegypti, for example, the expression levels of allatotropin receptor

vary between brain, gut and reproductive tissues and receptor levels fluctuate according to the

developmental and feeding state of the organism (Nouzova et al., 2012). Hence, a single

neuropeptide can conduct a coordinated physiological response by affecting multiple tissues

4

simultaneously. Moreover, multiplicity can be achieved by the presence of several isoforms of a

neuropeptide that are in turn released or expressed in a time and tissue-specific manner e.g.

allatotropins, allatostatins, leucokinins, adipokinetic hormones and tachykinins can all have

multiple isoforms within single species (Nässel, 2002). Additionally, multiple neuropeptides

may be co-localised and co-released, or released as sequential peptidergic ensembles from

neurosecretory cells, resulting in increased complexity and efficacy in regulation of

physiological processes. Examples of such coordinated biological behaviours include feeding,

ecdysis and reproduction (Audsley and Weaver, 2009; Kim et al., 2006b; Truman, 2005;

Wielendaele et al., 2013).

Familial Organization

Neuropeptide nomenclature stems from grouping peptides with similar biological functions or

sequence similarities under one family. However, sometimes, structurally unrelated peptides are

also grouped together based on their preliminary identification e.g. members of the allatostatin

family: allatostatin A, B and C. These allatostatins are derived from three different genes, have

different peptide sequences and not all have allatoregulatory function across species;

nevertheless they are collectively classified as allatostatins. Hence, while most neuropeptides

are organized into respective functional or structural families, some discrepancies do exist and

these will probably increase as additional (related or unrelated) functions of neuropeptides are

discovered.

Identification

Traditionally neuropeptides were identified via mass tissue extraction followed by peptide

sequencing through Edman degradation. Matrix-assisted laser desorption/ionization time-of-

flight mass spectrometry (MALDI-TOF MS) and immunohistochemistry with antibodies

specific to the peptide have aided in determining peptide sequence and distribution within

peripheral tissues and central nervous systems. However, with the recent availability of genomic

databases for various arthropod species, prediction and sequencing of genes encoding

neuropeptides and candidate receptors has become possible. Therefore, specialized techniques

such as in situ hybridisation, using transcript specific probes, can now be deployed to localize

neuropeptide gene transcripts in cells. Moreover, with molecular techniques such as qRT-PCR

5

one can quantify the neuropeptide transcript levels in various tissues at different developmental

time points. Additionally, specific biological roles of neuropeptides in a given process can be

assigned by altering neuropeptide expression levels via gene silencing through RNA

interference (RNAi) and GAL4/UAS systems. Receptor identification and characterisation has

further fuelled neuropeptidomic research and thus vital information about specific neuropeptide-

receptor interactions can be attained along with an insight into possible second messenger

pathways involved. Although genomic sequencing and annotation have accelerated

neuropeptide research, deficiencies still exist in our comprehension of post-translational

modifications and the structure of the mature neuropeptides eventually produced; hence, these

new methods of genetic sequencing and bioinformatics need to be complemented with

traditional techniques of mass spectrometry for determining the complete biochemical nature of

the mature neuropeptides (Caers et al., 2012; Nässel and Winther, 2010).

Synthesis and Release

Neuropeptides are secretory proteins transcribed with signal sequences guiding them to the

endoplasmic reticulum for post-translational modifications. They are synthesised as large

precursor molecules that can encode one or many structurally-related or unrelated mature

neuropeptides, each subsequently cleaved at specific dibasic cleavage sites (Caers et al., 2012;

Veenstra, 2000). Following post-translational modifications such as glycosylation, N-terminal

pyroglutamate addition, C-terminal amidation etc the neuropeptides are packaged into secretory

storage granules and channelled to axon terminals where they are stored and can undergo

exocytic release when the membrane is depolarized (Nässel, 2002; Zupanc, 1996).

G-protein coupled receptors (GPCRs)

G-protein coupled receptors (GPCRs), as the name suggests, are transmembrane receptors

associated with heterotrimeric GTP binding proteins, G proteins, at their cytosolic C-terminal

(Caers et al., 2012). The structure of GPCRs is conserved and defined by the presence of 7 α-

helical chains that span the membrane thus linking the outside of the cell to the inside. These

helical chains are referred to as the transmembrane domains (Meeusen et al., 2003). Messages

are transduced when specific ligands such as neuropeptides or biogenic amines (Hauser et al.,

6

2008) interact with sequences at the extracellular, usually glycosylated, N-terminus. Ligand-

binding initiates a cascade of intracellular events that begin with activation and dissociation of

the heterotrimeric GTP molecule via dephosphorylation. The dissociated units subsequently

interact with various host molecules, such as phospholipase Cβ or adenyl cyclase, resulting in

either an increase of intracellular Ca2+

levels or synthesis of 3’-5’-cyclic adenosine

monophosphate (cAMP). Moreover, the subunits can interact directly with ion channels; Ca2+

or

K+

ion channels, resulting in an influx of extracellular calcium and potassium (Caers et al.,

2012; Meeusen et al., 2003). Receptor activation by certain ligands can be determined by

measuring these intracellular Ca2+

or cAMP levels with the help of cell lines cells expressing

apoaequorin proteins or the luciferase gene respectively (Caers et al., 2012; Hauser et al., 2006).

Additionally, receptor activation can also be used to identify receptor-specific ligands from a

library of neuropeptides, and hence isolated, putative receptors can be deorphanized (reverse

pharmacology). The entire signal transduction pathway culminates in ligand degradation or

receptor desensitization/internalization due to prolonged stimulation. Receptor-ligand

interactions are an important avenue to explore due to their manifestation as mediators of critical

biological roles such as reproduction, feeding, growth and behaviour. Biosynthetic agents

engineered towards altering such interactions could be pivotal in designing species-specific

pesticides.

Allatoregulatory Peptides

Allatoregulatory peptides are a class of neuropeptides that have been identified as regulators of

JH biosynthesis from the CA of insects. These include stimulatory allatotropins and inhibitory

allatostatins. Juvenile hormones are sesquesterpenoids produced by the endocrine CA

(Goodman et al, 2005) that govern key biological processes in insects including development

and metamorphosis (Tobe and Bendena, 1999), reproduction including vitellogenesis and egg

production (Davey, 2007), and behaviour (Nijhout and Wheeler, 1982; Riddiford, 2008; Zhu et

al., 2008). Evolutionary analyses reveal ‘acquired roles’ for JH regulation in developmental

processes (ecdysis and metamorphosis) and subsequent modulation of this hormone by

allatoregulatory peptides (Tobe and Bendena, 1999). Hence, despite the recent revelation of

stage, species and dose-dependent regulation of allatoregulatory peptides on JH titres (Audsley

7

et al., 2008; Hoffmann et al., 1999; Weaver and Audsley, 2009), the possible additional

ancestral roles that may still be modulated by these allatoregulatory neuropeptides must also be

taken into consideration. Figure 1 represents a schematic adapted from McNeil and Tobe (2001)

that summarises some of the main physiological roles regulated by allatoregulatory peptides;

however, it must be noted that allatoregulatory peptides are pleiotropic in nature and only a

subset from the plethora of physiological functions mediated by these neuropeptides has been

represented in Figure 1.

Allatostatins

Allatostatins (ASTs) belong to a family of structurally-unrelated peptides that were originally

isolated as inhibitory neuropeptides affecting JH biosynthesis (Nässel, 2002). Due to their

structural disparity ASTs have been grouped into three distinct families: allatostatin-A (AST-A),

allatostatin-B (AST-B) and allatostatin-C (AST-C); each family having been identified from

different species and each possessing multimodal characteristics. The AST-A, also called the

FGLamides due to their characteristic C terminal Y/FXFGL/I/V sequence and AST-C, also

known as PISCF-ASTs due to their C terminal motif, were originally isolated as inhibitors of JH

from Diploptera unipuncta and Manduca sexta respectively (Kramer et al., 1991; Woodhead et

al., 1989). Based on these findings, they are alternatively also referred to as the Diploptera type

or Manduca sexta type ASTs; however, since then both AST-A and AST-C have been identified

in numerous other species where they are found to express allatostatic properties along with

inhibitory roles in a variety of physiological processes e.g. feeding (Weaver and Audsley, 2009).

In addition, AST-B type neuropeptides (W2W

9) were first isolated as myoinhibitory peptides

(MIPs) from L. migratoria (Nässel, 2002; Schoofs et al., 1991) and later identified as inhibitors

of JH biosynthesis in M. sexta and G. bimaculatus (Nässel, 2002); hence, the AST-B family is

also acknowledged as Lom-MIP or locust-type AST. Therefore, it is evident that allatostatins

exert a species specific inhibition on JH biosynthesis and are not universal in their allatostatic

function as initially considered.

8

Figure 1. A schematic representing the possible targets of allatoregulatory peptides (AT and

AST). Modified from McNeil and Tobe (2001).

9

Allatotropins

Identification and Characterization

Another set of neuropeptides that regulate JH biosynthesis from the CA are the family of

allatotropins. Allatotropins (ATs) were first isolated as amidated tridecapeptides from the heads

of Manduca sexta pharate adults and subsequently shown to have stimulatory effects on JH

biosynthesis from the CA (Katoaka et.al., 1989). Since then peptides similar to AT have been

characterised or predicted in various holometabolous and hemimetabolous insects (Table 1).

Surprisingly though, ATs are also present in non-insect species such as arachnids (Horodyski,

2013), annelids (Ukena et al., 1995), crustaceans (Christie et al., 2011) and even molluscs;

species that lack JH (Jing et al., 2010; Veenstra, 2010).

The mature, functional peptide is usually 13-14 residues long, with a highly conserved

pentapeptide TARGF-amide C-terminal motif which constitutes the biologically-active domain

(Table 1; Audsley et al., 2008). Hitherto, only two structurally-unrelated ATs have been

identified; the classic TARGF-amide AT and AT-2, a unique allatotropin isolated from S.

frugiperda with a C-terminal PISCF sequence (Abdel-latief et al., 2004a). AT-2 has a C-

terminal sequence similarity with the traditional AST-C and was expected to be allatostatic in

nature; however, due to its stimulatory effect on JH biosynthesis it was termed AT-2 (Abdel-

latief et al., 2004a). To date, this novel AT is limited to S. frugiperda and its presence has not

been identified in any other organism.

Allatotropin and Juvenile Hormone

Allatotropins, as implied by their name, are potent stimulators of JH biosynthesis from the CA

of some insects. They were first identified as allatoregulatory peptides in adult M. sexta were

they were able to induce an 80% increase in JH biosynthesis. Subsequently, ATs have been

found to regulate JH titres in a dose–dependent and stage specific manner in several insect

species including lepidopterans: S. frugiperda, L. oleracea, H. virescens, (Audsley et al., 2000;

Oeh et al., 2000; Rachinsky et al., 2003); dipterans: A. aegypti and P. regina (Li et al., 2003; Tu

et al., 2001), a hymenopteran: A. mellifera (Rachinsky and Feldlaufer, 2000) and a

dermapteran: E. annulipes (Rankin et al., 2005). It has been confirmed by in-vitro and receptor-

10

Table 1. Sequences of known or predicted allatotropin peptides. Amino acids shared by all

sequences are highlighted in black, while conserved amino acids are shaded grey.

Species Peptide Sequence References

Lepidopterans Manduca sexta GFK-NVEMMTARGF (Taylor et al., 1996)

Bombyx mori GFK-NVEMMTARGF (Nagata et al., 2012)

Spodoptera frugiperda GFK-NVEMMTARGF (Abdel-latief et al., 2003)

Samia cynthia ricini GFK-NVEMMTARGF (Sheng et al., 2007)

Helicoverpa armigera GFK-NVEMMTARGF (Yin et al., 2005)

Pseudaletia unipuncta GFK-NVEMMTARGF (Truesdell et al., 2000)

Diptera

Aedes aegypti APFR-NSEMMTARGF (Veenstra and Costes, 1999)

Anopheles gambiea APFR-NSEMMTARGF (Riehle et al., 2002)

Hemiptera

Rhodnius prolixus GFK-NVQLSTARGF

(Ons et al., 2009; Ons et al.,

2011)

Ixodida

Ixodes scapularis GFR-KMKISTARGF (Horodyski, 2013)

Hymenopetra

Nasonia vitripennis GFQ-PEYISTAYGF (Hauser et al., 2010)

Cladocera

Daphnia pulex GFK-TVGLATARGF (Christie et al., 2011)

Coleoptera

Tribolium castaneum GIEALKYH-NMDLGTARGY (Weaver and Audsley, 2008)

Orthoptera

Locusta migratoria GFK-NVALSTARGF (Paemen et al., 1991)

Blattodea

Periplaneta americana GFK-NVALSTARGF (Neupert et al., 2009b)

Leucophaea maderae GFK-NVALSTARGF (Neupert et al., 2009b)

Mollusca

Aplysia californica GFRLNSASRVAHGY (Jing et al., 2010)

Lottia gigantea GFKANSASRVAHGY (Veenstra, 2010)

Haplotaxida

Eisenia foetida GFKDGAADRISHGF (Elekonich and Horodyski,

2003; Ukena et al., 1995)

11

mediated assays that the mode of action of AT upon JH biosynthesis stimulation is through Ca2+

and cAMP dependent-pathways, whereby an increase in the intracellular Ca2+

levels of the CA

cells results in an increased JH production (Horodyski et al., 2011; Rachinsky et al., 2003;

Vuerinckx et al., 2011).

Multimodal Roles of Allatotropin

Allatotropins are pleiotropic peptides that have been found to regulate diverse physiological

processes and are hence not just limited to stimulation of JH biosynthesis from the CA of

insects. Veenstra (1994) identified a cardioacceleratory role of AT in the tobacco hornworm, M.

sexta, and since then ATs have been found as cardioacceleratory peptides stimulating

contractions in a dose-dependent manner in various species including L. maderae, P. americana

(Rudwall et al., 2000) and T. infestans (Sterkel et al., 2010). Moreover, ATs have also been

identified as myostimulatory peptides inducing gut contractions (foregut, whole gut or hindgut)

in multiple lepidopteran and blattodea species, along with a coleopteran, T.castaneum,

dermapteran: E. annulipes and hemipteran: T. infestans (Duve et al., 1999; Oeh et al., 2003;

Rankin et al., 2005; Santini and Ronderos, 2007; Vuerinckx et al., 2011). Additionally, in M.

sexta ATs have been found to be potent inhibitors of ion transport across the posterior midgut

epithelium; however, this is a stage and species-specific response that has only been observed in

feeding fourth instars and day 2 fifth instars, and is completely absent in other lepidopterans

(Lee et al., 1998).

Allatotropin-related peptides (ATRPs) have also been identified from the male accessory glands

of L. migratoria (Lom-AG-MT) and head extracts of Leptinotarsa decemlineata, (Led-OVM)

(Paemen et al., 1991; Spittaels et al., 1996). These ATRPs are found to have a preferential

myostimulatory effect on female oviducts than hindguts of both aforementioned species as well

as Leucophaea maderae. Myotropic regulation of reproductive tissues is another novel role of

ATs that opens avenues for research into other reproductive processes (vitellogenesis, sexual

receptivity, oviposition rate, egg deposition) that are modulated by AT either directly,

myostimulation, or indirectly via JH regulation; a role similar to the sex peptides of Drosophila

(Chen et al., 1988; Elekonich and Horodyski, 2003).

12

A possible role in photic entrainment of the circadian clock has also been suggested for AT. In

monarch butterflies, JH levels are up-regulated in summer butterflies compared to migratory

butterflies. The increased JH levels in suitable environmental conditions allow for rapid

completion of female ovarian cycle, vitellogenesis, as well as production of sex pheromones -

all factors leading to an onset of sexual maturity in recently emerged females. Concurrently AT

levels are also up-regulated in summer monarch butterflies vs migratory butterflies suggesting a

possible allatoregulatory role of the neuropeptide upon JH production (Zhu et al., 2008). Similar

JH up-regulation was also observed in summer vs migratory individuals of the true armyworm,

P. unipuncta (McNeil and Tobe, 2001); however, since AT levels were not quantified in the two

behavioural phenotypes, it is only speculated that a similar JH stimulation and up-regulation of

AT levels occurs within the sedentary summer moths. Moreover, AT has been found to be

associated with the circadian rhythms of L. maderae, where time-dependent injections cause a

phase shift in the circadian rhythms of the innate locomotor activity (Petri et al., 2002; Petri et

al., 1995).

The presence of ATs in non-insect phyla such as annelids and molluscs, species that lack JH,

suggests an alternative role for AT. Indeed, it has been found that the allatotropin-related

peptide present in Aplysia regulates muscle contractions in the protraction behaviour (Jing et al.,

2010). Moreover, in the Turbellaria and Catenulida AT-like immunoreactivity has been found

associated with muscle tissues suggesting a possible myotropic activity in these taxa (Adami et

al., 2011). Considering these findings, it has been suggested that the ancestral role of AT is

myotropic, with allatoregulation of JH titres being an acquired function in arthropods over time

(Elekonich and Horodyski, 2003; Tobe and Bendena, 1999).

Allatotropin Gene

The family of ATs, including allatotropin-like (ATL) and allatotropin related peptides (ATRPs),

have been identified and extensively characterised among the invertebrate phyla including

insects, molluscs, annelids, arachnids and crustaceans (Christie et al., 2011; Jing et al., 2010;

Ukena et al., 1995; Veenstra, 2010); however, genomic characterisation has been largely limited

to lepidopterans (Abdel-latief et al., 2003; Nagata et al., 2012; Sheng et al., 2007; Taylor et al.,

1996; Truesdell et al., 2000; Yin et al., 2005) with the only species outside this order from

which the AT gene has been isolated and cloned being the dipteran, A. aegypti (Veenstra and

13

Costes, 1999). The AT neuropeptide is encoded by a single gene as a single transcript, except in

lepidopterans where multiple splice variants are present. The number of variants and hence the

number of isoforms expressing AT and ATL peptides are species-specific and range from 3-4

isoforms per species. Though ATL peptides do not share a sequence homology to AT, they are

relatively well conserved amongst themselves (Table 2). Allatotropin and ATL peptides are

preferentially expressed in certain tissues of the organism in a time- and developmental-stage

specific manner (Horodyski et al., 2001) thereby adding to the multiplicity and hormonal

complexity of allatotropin and its functions. Besides the lepidopterans, the only other species

predicted to have multiple isoforms is Ixodes scapularis (Horodyski, 2013). With the

commencement of the peptidomic era, ATs have also been predicted or determined in other

insects (Table 1) using in silico bioinformatic tools to screen the genomes and/or mass

spectrometric analyses. Initially thought to be absent from the parasitic wasp: Nasonia

vitrepennis (Hauser et al., 2010), a subsequent in silico genomic analysis has predicted the

presence of a hypothetical AT protein (Ac: XP_003427574.1); however, despite the presence of

this genomic toolkit, no AT or ATRPs have been identified in Drosophila species (Adams et al.,

2000; Nässel, 2002; Nässel and Winther, 2010).

Allatotropin Receptor

The allatotropin receptor (ATR) has been identified from the yellow fever mosquito, A. aegypti

(Nouzova et al., 2012); Tribolium castaneum (Vuerinckx et al., 2011); and two lepidopterans:

Bombyx mori (Yamanaka et al., 2008) and M. sexta (Horodyski et al., 2011). The receptors

share a high sequence similarity with mammalian orexin receptors, suggesting a possible

evolutionary link between the two (Caers et al., 2012). Analysis of the Drosophila genome

again yielded no results for the presence of ATRs (Caers et al., 2012).

14

Table 2. Sequence similarities, highlighted in black, between allatotropin-like peptides of

different lepidopteran species

Species Sequence References

Manduca sexta_I GTFKPNSNILIARGY (Horodyski et al., 2001)

Samia cynthia ricini_I GTFKPNSNILIARGY (Sheng et al., 2007)

Helicoverpa armigera_I GVFRPNSNVLIARGY (Yin et al., 2005)

Manduca sexta_II GTPTFKSPTVGIARDF (Horodyski et al., 2001)

Samia cynthia ricini_II GTPIFKSPTVGIARDF (Sheng et al., 2007)

Helicoverpa armigera_II GTPTFKSPTVGIARDF (Yin et al., 2005)

Manduca sexta_III PWFNPKSKLLVSTRF (Horodyski et al., 2001)

Samia cynthia ricini_III FNPKNSLMVAYDF (Sheng et al., 2007)

Helicoverpa armigera_III FNPKSNLMVAYDF (Yin et al., 2005)

15

Allatotropin receptors are expressed in a tissue specific manner with high transcript levels in the

central nervous systems and reproductive tissues, with the only exception of high transcript

levels in the guts of M. sexta. The receptor transcript exhibits sex specific expression in T.

castaneum (Vuerinckx et al., 2011), while in the lepidopterans and dipterans, transcript levels

are regulated according to the developmental stage and feeding state of the organism e.g. a

decrease in receptor expression is observed in A. aegypti following a blood meal (Nouzova et

al., 2012). The receptor expression levels follow endogenous JH titres through different

developmental stages. In lepidopterans, allatotropin receptors exhibit a preferential binding to

AT and ATL peptides in a dose-dependent manner, whereby ATL-I peptide, with a semi-

conserved C-terminal domain, is more potent than AT itself (Horodyski et al., 2011). The ATRs

are typical GPCRs that activate the phospholipase Cβ and adenyl cyclase pathway leading to an

increase in intracellular Ca2+

levels and cAMP levels. It has been found in H. virescens that an

increased Ca2+

concentration leads to an increased sensitivity of the CA cells (Rachinsky et al.,

2003). Moreover, in the same year Oeh et al. (2003) discovered that the signal transduction

pathway for the myostimulatory effects of AT on visceral muscles of H. virescens was also

mediated by an increase in intracellular Ca2+

levels through the PKC pathway. Hence, signal

transduction through Ca2+

mediated pathways seems to be the primary route of action for ATs.

Allatotropins and Rhodnius prolixus

In 2009, Ons et al. isolated three putative, truncated isoforms of AT from R. prolixus brains

following a neuropeptidomic search using mass spectrometry. In a subsequent study, however,

Sterkel et al. (2011) determined that the truncated isoforms were probably structurally-

redundant and only one mature amidated tridecapeptide was thought to be the functional AT

(Rhopr-AT). Subsequent research by Ons et al. (2011) led to the discovery, isolation and

characterisation of the gene encoding AT in R. prolixus; however, these few aforementioned

experiments encapsulate the extent of our knowledge regarding ATs in R. prolixus, and aspects

such as gene expression, distribution and peptide function are still untouched.

Since some research has been done in a closely related triatomine bug, Triatoma infestans, an

initial comparative approach can be taken to study the distribution and roles of AT in R.

prolixus. In T. infestans, allatotropin-like immunoreactive (AT-like-ir) processes have been

observed extending over the dorsal vessel and retrocerebral complex; while AT-like-ir cells are

16

also seen in the optic lobes of the brain and in the anterior midgut as open type endocrine cells

(Riccillo and Ronderos, 2010). Surprisingly, however, AT-like immunoreactivity is also

observed in the Malpighian tubules (MTs) and the AT-like-ir content is regulated depending

upon the feeding state of the insect (Santini and Ronderos, 2009). T. infestans, like R. prolixus,

are blood gorging hemipterans that imbibe a large blood meal and are hence faced with the same

short-term physiological challenges as R. prolixus; they need to rapidly undergo diuresis. Hence,

based on the hypothesis that AT produced from the endocrine cells of the MTs regulates diuretic

behaviour it was found that AT exerted a myotropic effect on hindgut contractions in a dose-

dependent manner (Santini and Ronderos, 2007). Moreover, AT was a sexually dimorphic

cardioacceleratory peptide at the level of the dorsal vessel and myostimulatory peptide upon the

anterior midgut (Sterkel et al., 2010).

THESIS OBJECTIVE

The goal of this thesis was to identify and confirm the gene transcript, its relative tissue

distribution and to determine any potential roles of AT in the blood-gorging hempiteran, R.

prolixus. These objectives were achieved through in silico bioinformatics, in vitro molecular

techniques supplemented with immunohistochemical analysis, and tissue bioassays. Moreover,

much of the work stemmed from comparative analysis with previous literature that determined

novel and peculiar roles of AT in T. infestans.

Originally isolated as stimulators of JH, ATs have since been identified as multimodal

neuropeptides that are well-represented among invertebrates. Allatotropins regulate a plethora of

physiological functions in complex combinations of species, tissue, sex or developmental time,

hence offering great complexity and multiplicity in function. Therefore, owing to its

fundamental role in an array of biological process it becomes critical to obtain a comprehensive

understanding of this family of neuropeptides that were once thought to be mere stimulators of

JH biosynthesis.

17

MATERIALS AND METHODS

Animals and tissues

Rhodnius prolixus were obtained from a long-standing colony maintained at high relative

humidity in 25oC incubators at the University of Toronto Mississauga. Insects were fed

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

stages. The blood meal initiates growth and development and the insect emerges into the next

instar approximately 1-2 weeks post-feeding. Unfed (approx. 4-6 weeks) IVth

instars were used

for conducting all the experiments with the exception of hindgut assays where both unfed IVth

and Vth instars were used. Additionally, the heart assays and RT-PCRs were conducted using

only unfed Vth

instars. All physiological assays were carried out in R. prolixus saline modified

from Lane et al (1975); namely150mM NaCl, 8.6mM KCl, 2.0mM CaCl2, 4.0mM NaHCO3,

8.5mM MgCl2, 5mM HEPES and 34mM glucose; pH 7.0. Tissues for molecular biology

experiments were dissected in nuclease free phosphate buffered saline (PBS) (Sigma-Aldrich,

Oakville, ON, Canada) and immediately placed in RNAse-free microcentrifuge tubes containing

RNA laterTM

RNA stabilization reagent (Qiagen Inc., Mississauga, ON, Canada).

Construction of CNS cDNA

Fresh IVth

instar central nervous system (CNS) cDNA was used as a template for PCR reactions.

Total CNS RNA was extracted using Trizol® reagent (Life Technologies Corporation,

Carlsbad, CA, USA) and 400ng of extracted RNA was used for cDNA synthesis using the

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

Ltd., Mississauga, ON, Canada). The cDNA synthesis reaction was carried out using the

following thermal cycler conditions: incubation at 25oC for 5 minutes, 42

oC for 30 minutes and

final 85°C for 5 minutes to heat-inactivate the reverse transcriptase. The final product was

diluted 1:2 and stored at -20oC, an aliquot being used for subsequent PCR reactions as needed.

18

In-silico and in-vitro search of cDNA sequence encoding allatotropin prepropeptide

Predicted allatotropin prepropeptide sequence (Ons et al., 2011) was used to tBLASTn search

the Rhodnius genome using Geneious software 4.7.6 (Biomatters, Ltd., Aukland, New Zealand).

The tBLASTn search yielded positive hits against contig 3631 and contig 64. The results

obtained were then used to predict possible promoter regions and splice sites via online software

provided by Berkeley Drosophila Genome Project. The possible signal peptide sequence was

predicted using SignalP 4.1 software (Petersen et al., 2011).

IVth

instar CNS cDNA was screened for R. prolixus allatotropin (Rhopr-AT) gene using gene

specific forward and reverse primers: Allatrp-For1, Allatrp-Rev5 (Table 3) in a Polymerase

Chain Reaction (PCR). These primers successfully amplified the entire open reading frame

(ORF) as well as segments of the 5’ and 3’ untranslated regions (UTRs). All PCRs were

conducted using the s1000 Thermal Cycler (Bio-Rad Laboratories, Mississauga, ON, Canada),

and the temperature cycling conditions were as follows: initial denaturation at 94oC for 3

minutes, followed by 35 cycles of denaturation at 94oC for 30 seconds, annealing at 62

oC for 30

seconds, extension at 72o C for 1 minute, followed by 10 minute final extension at 72

o C.

Products obtained from the PCR were column purified using the AxyPrep™ PCR Clean-up Kit

(Axygen Biosciences, Union City, CA, USA) and subsequently cloned into the pGEM®-T

vector (Promega Corporation, Madison, WI, USA). Mach1™-T1R Chemically Competent E.

coli cells (Life Technologies Corporation, Carlsbad, CA, USA) were transformed with the

recombinant vector and grown on LB/Ampicillin/XGal medium for 12-16 hours. Successfully

transformed colonies were selected; their plasmids purified and sent for sequencing at the Sick

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

Toronto, ON, Canada).

The molecular techniques remained identical for the cloning of PCR products that were

subsequently used for synthesis of RNA probes for Northern blot hybridisation, except for gene

specific primers, annealing and extension temperatures and times. The primers used for

amplification of this product were Allatrp-For1-Exon3 and Allatrp-Rev5 at an annealing

temperature of 51oC for 30 seconds and an extension step at 72

oC for 30 seconds. The rest of the

thermal cycler conditions remained unchanged.

19

Table 3. Gene-specific primers used for Rhopr-AT prepropeptide cDNA amplification

Oligonucleotide Name Oligonucleotide sequence (5’-3’)

Allatrp-For1 GCATCAGTCGGAGTACAGA

Allatrp-For1-Exon3 TGTCCATCGTCAGCATAC

Allatrp-Rev4 AGTATGCTGACGATGGACAC

Allatrp-Rev5 CAGATGAAATGACTTTTTAACAACAC

rp49for1 (positive control) GTGAAACTCAGGAGAAATTGGC

rp49Rev1(positive control) AGGACACACCATGCGCTATC

20

Phylogenetic Analysis of Rhopr-AT prepropeptide

In order to identify homologous allatotropin sequences from other insect species a protein

BLAST against other genomic databases was performed using Rhopr-AT prepropeptide

sequence (Ac: ACS45387.1) as a query. The prepropeptide sequences were selected from the

following species: Manduca sexta (Ac: AAB08757.1), Mythimna separata (Ac: ABA61322.1),

Mythimna unipuncta (Ac: AAF23016.1), Spodoptera frugiperda (Ac: CAD32495.1),

Spodoptera exigua (Ac: AEO27700.1), Helicoverpa armigera (Ac: AAR32789.1), Samia ricini

(Ac: AAZ30065.1), Daphnia pulex (Ac: EFX71302.1), Clostera anastomosis (Ac:

AAV59461.2), Aedes aegypti (Ac: AAB06179.1), Anopheles gambiae (Ac: Xm_320402.3),

Bombyx mori (Ac: AAY40838.1), Tribolium castaneum (Ac: NP_001137204.1), Ixodes

scapularis (Ac: EEC06620.1), predicted hypothetical protein: Nasonia vitripennis (Ac:

XP_003427574.1), predicted hypothetical protein: Culex quinquefasciatus (Ac: EDS30439.1),

predicted hypothetical protein: Acyrthosiphon pisum (Ac: XP_001942912.1), predicted

hypothetical protein: Bombus impatiens (Ac: XP_003488345.1) and predicted hypothetical

protein: Bombus terrestris (Ac: XP_003398476.1). Aforementioned sequences were used to

conduct a subsequent alignment using ClustalOmega (http://www.ebi.ac.uk/Tools/msa/clustalo/)

following which, the sequences were imported to BOXSHADE

(http://www.ch.embnet.org/software/BOX_form.html) to construct the final image depicting

conserved regions between amino acid sequences. The sequences along with splice variants, if

any, were imported into MEGA 5.0 (Tamura et al., 2011) and a maximum parsimony

phylogenetic tree was constructed using the Close-Neighbor-Interchange (CNI) analysis with a

bootstrap value of 1000 iterations.

Reverse Transcription-Polymerase Chain Reaction (RT-PCR) to elucidate spatial distribution

of transcript

Insects were dissected and the tissues pooled into 11 separate groups: 1) CNS, 2) dorsal vessel

(DV), 3) foregut (FG), 4) salivary glands (SG), 5) fat body and trachea (FbTr), 6) anterior

midgut (AM), 7) posterior midgut (PM), 8) Malpighian tubules (MT), 9) hindgut (HG), 10)

immature testes (TT) and 11) immature ovaries (Ov). Total RNA from these tissues was

extracted using the Trizol® reagent (Life Technologies Corporation, Carlsbad, CA, USA) and

http://www.ebi.ac.uk/Tools/msa/clustalo/http://www.ch.embnet.org/software/BOX_form.html

21

cDNA was synthesised as described earlier. To elucidate the presence of Rhopr-AT mRNA

transcript a 553bp fragment spanning all three exons was chosen. This region was amplified

using the following primers: Allatrp-For1 and Allatrp-Rev4, while a positive control with the

amplification of a 229bp ribosomal protein 49 (Rp49) was done using rp49for1 and rp49Rev1

primers (Table 3). The thermal cycle conditions for the ensuing PCR reaction were as follows:

initial denaturation at 94°C for 3 minutes, 35 cycles of denaturation at 94°C for 30 seconds,

annealing at 57°C for 30 seconds, extension at 72°C for 30 seconds and final extension at 72°C

for 10 minutes. Three independent biological replicates were done for the RT-PCR.

Northern Blotting to determine transcript size

Total CNS RNA from 100 insects was extracted using Trizol® Reagent (Life Technologies

Corporation, Carlsbad, CA, USA). 2µg of RNA along with 2x RNA loading dye (Fermentas

Canada Inc., Burlington, ON, Canada) was denatured at 75oC for 5 minutes and immediately

chilled on ice. 30ng of RiboRuler™ High Range RNA Ladder (Fermentas Canada Inc.,

Burlington, ON, Canada) with 2x RNA loading dye (Fermentas Canada Inc., Burlington, ON,

Canada) was also prepared under simultaneous conditions. Samples were loaded on a 1% RNase

free agarose gel containing formaldehyde and electrophoresed at 70V for 140 minutes. To

ensure the RNA samples were separated sufficiently the gel was briefly viewed under UV light.

Following UV analysis, a 20 minute wash with diethylpyrocarbonate (DEPC)-treated distilled

water was done, and the RNA was transferred from the gel onto a positively charged nylon

membrane (Roche, Mannheim, Germany) in 20x saline-sodium citrate (SSC) via overnight

downward capillary transfer. The gel was briefly viewed under UV light again to ensure no

residual RNA remained. The nylon membrane was briefly washed (4-5 seconds) in DEPC

treated water to remove excess 20xSSC that could result in high background of the blot. Finally,

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

Upland, CA) and drying the membrane at room temperature for 2-3 hours. The blot was stored

at 4oC until further use.

Hybridization was conducted using the DIG Northern Starter Kit (Roche, Mannheim, Germany)

with some modifications to the manufacturer’s protocol. Specifically, a digoxigenin (DIG)-

labelled anti-sense RNA probe complementary to the 3’ UTR of Rhopr-AT mRNA transcript

was synthesised for hybridisation with the membrane. Overnight linearization at 37oC of a 20µg

22

pGEM®-T vector, containing 426bp Rhopr-AT fragment, was done using SacII Restriction

Enzyme. The reaction was then quenched with a water bath treatment at 65oC for 20 minutes

and the linearized plasmid was PCR purified using the AxyPrep™ PCR Clean-up Kit (Axygen

Biosciences, Union City, CA, USA). In vitro transcription of the 1µg of linearized plasmid

using Sp6 polymerase from the DIG RNA labelling kit SP6/T7 (Roche Applied Science,

Mannheim, Germany) followed. The DNA template was removed by incubating the reaction

with 2µl of deoxyribonuclease I for 15 minutes at 37oC. The synthesised antisense RNA probe

was stored at -20oC until further use. A DNA DIG-labelled RNA ladder probe was also

synthesised according to the manufacturer’s protocol and stored at -20oC.

The RNA-bound nylon membrane was pre-hybridized for 30 minutes at 68oC with pre-warmed

hybridisation solution. Following pre-hybridisation, the membrane was hybridised for 16-18

hours at 68oC with hybridisation solution containing100ng/mL SacII linearized RNA probe. To

remove unbound probe and decrease background noise, the membrane was subjected to

stringent washes: two washes with 2x SSC, 0.1% SDS at room temperature, followed by two

washes with pre-warmed 0.5x SSC, 0.1% SDS at 68oC. Immunological detection of the

membrane was done by exposing the Bioflex Scientific Imaging Films (Clonex Corporation,

Markham, ON, Canada) to the chemiluminescence substance 15-30 minutes post application.

Transcript size of the Rhopr-AT prepropeptide was determined by aligning it to the RNA ladder.

Three independent biological replicates were performed.

Immunohistochemistry to detect peptide localisation

The dorsal cuticle and diaphragm of unfed IVth

instar R. prolixus nymphs were removed to

expose the digestive tract and CNS. Tissues were subsequently fixed in-situ by submerging the

bug in 2% paraformaldehyde fixative (pH 7.0) for 18-24 hours at 4˚C. Excess fixative was

washed off using phosphate buffered saline (PBS) - washes were done every 10 minutes for 2

hours. The tissues (gut, Malpighian tubules, salivary glands and CNS along with peripheral and

abdominal nerves) were dissected out and placed into respective 1.5 ml microcentrifuge tubes

containing PBS. To increase permeability of tissues to the antiserum they were treated with 4%

Triton X-100 in PBS with 2% bovine serum albumin (BSA) and 10% normal goat serum (NGS)

23

for 1 h at room temperature. Excess Triton X-100 was washed off with PBS and the tissues were

incubated on a shaker for 48 hours at 4oC with antiserum made by rabbits against Manduca

sexta AT (Manse-AT: GFKNVEMMTARGF-NH2, a gift from J. Veenstra, France). The

antiserum had been pre-incubated for 18-24 hours at 4oC with a 0.4% solution of Triton X-100

in PBS with 2% BSA and 2% NGS to obtain a 1:1000 dilution and to minimise non-specific

binding. Following primary antiserum incubation, tissues were washed at room temperature for

4-6 hours in PBS and then incubated on a shaker for 18 hours at 4oC with a secondary antibody

of Cy3-labelled goat anti-rabbit diluted 1:600 with 10% NGS in PBS. Tissues were then washed

with PBS repeatedly for 8 hours and finally mounted on microscope slides in 100% glycerol.

Allatotropin-like immunoreactivity was observed and images were obtained using a Zeiss

confocal laser microscope (Carl Zeiss, Jena, Germany) and Zen 2009 viewing software.

Allatotropin-like immunoreactivity within the CNS was mapped using an epifluorescence

microscope with a drawing tube attached. Controls whereby primary antiserum was pre-

incubated with 10-5

M Rhopr-AT for 24 hours yielded no staining.

Physiological Roles of Allatotropin: Hindgut Contraction Assay

A horizontal incision along the thoracic segment and two lateral incisions along the dorsal

cuticle of R. prolixus were done to expose the tissues. Tissues were bathed in 100µl R. prolixus

saline. Dorsal diaphragm, dorsal vessel, Malpighian tubules, fat body and trachea were removed

sequentially in an effort to obtain a clean preparation consisting of just the gut. Then, a silk

thread was used to tie a double knot at the anterior end of the hindgut (at the junction of

posterior midgut and hindgut) and the rest of the anterior digestive tract removed. The hindgut,

attached to the ventral cuticle at the anus, was removed and pinned in a well containing 200µl of

saline in a Sylgard-coated Petri dish. The free end of the silk thread was used to tie a double

knot around the hook of the force transducer (Aksjeselskapet Mikroelektronikk, Horten,

Norway). The knots and the resulting thread were tied at a 90o angle with the hook to increase

efficiency of the force transducer. The hindgut was stretched slightly to mimic its natural

physiological state. Longitudinal contractions exhibited by the tissue were measured using

PicoScope 2202 and recorded with PicoLog (Pico Technology, Cambridgeshire, UK). The

hindgut was washed with saline and left to bathe and equilibrate in saline for about 20 minutes

24

prior to starting the experiment. Synthetic Rhopr-allatotropin (Table 4) was stored at -20 oC as

5µl aliquots of 10-3

M and serially diluted with saline to obtain the desired applied dose for the

tissue. All doses were made and kept on ice at the beginning of the experiment, and brought

back to room temperature prior to application. Allatotropin doses ranged from 1x 10-16

M to

1x10-6

M and were applied in a random order. For application of a dose, 100µl of saline was

removed and 100µl of saline solution containing twice the desired concentration of peptide was

added simultaneously to maintain the total volume of 200µl in the well. The change in basal

tonus from saline incubation to peptide application was measured and then the dose was washed

off with saline until the basal tonus returned to pre-dose conditions. The experiment was

repeated with at least 5 independent hindgut preparations. Data were analysed using GraphPad

Prism.

To verify the viability of the preparation and to determine the relative effects of allatotropin

compared to a known stimulator of hindgut contractions, Rhopr-kinin 2 (Table 4) (dose range:

10-11

M to 10-6

M) was also applied on the same prep. Furthermore, the same doses of Rhopr-AT

and Rhopr-kinin were also tested on Vth

instar hindguts to deduce if any differences in effect

existed between the two instar stages.

25

Table 4. Amino acid sequence and purity of peptides used for the bioassays

Peptide Name Peptide Sequence Peptide Purity

Rhopr-AT NVQLSTARGF-amide >98%

Rhopr-Kinin 2 AKFSSWG-amide >95%

Proctolin RYLPT >83%

26

Physiological Roles of Allatotropin: Heart Impedance Assay

The legs were removed and the bug was pinned dorsal side down in Sylgard-coated Petri dish.

Lateral incisions along the ventral cuticle and a single horizontal incision along the ventral

thoracic segment enabled smooth removal of the ventral cuticle. Following this, the ventral

diaphragm and digestive tract were also removed to expose the dorsal vessel and associated

alary muscles which were attached to the dorsal cuticle and diaphragm. Tissues were bathed in

200µl of saline. The experimental set up consisted of minuten pin electrodes on either side of

the heart at the junction of the 5th

and 6th

abdominal segments. The electrodes measured the

change in resistivity caused by every beat of the heart. This change in resistivity resulting in

impedance of electric flow between the electrodes was amplified using an impedance converter

(UFI model 2991, Morro Bay, CA, USA), measured with a PicoScope 2202 and recorded with

PicoLog Software (Pico Technology, Cambridgeshire, UK). Since the change in impedance is

due to the heart beat, every deflection upon the PicoScope chart is indicative of a heartbeat. The

heart was allowed to acclimatize to the experimental set up for about 15 minutes before starting

the experiment. Allatotropin doses (10-12

to 10-6

M) were made as described for the hindgut assay

and tubes were kept on ice until application upon the preparation. Doses were applied in a

random order, removing and adding the same volumes as mentioned earlier for the hindgut

assays. Positive controls using the myotropic pentapeptide, proctolin (Table 4) (10-10

and 10-9

M)

were also done to confirm viability of the preparation.

27

RESULTS

Characterisation of cDNA and determination of Rhopr-AT transcript length

An in-silico BLAST search followed by cloning of the Rhopr-AT gene using gene specific

primers revealed a cDNA region of 973bp (Figure 2A). The cloned cDNA was identical to the

predicted sequence by Ons et al. (2011; Ac: GQ162783). The cDNA spans over three exons:

83bp, 211bp and 680bp in length. The first exon, situated on contig 3631 contains the

transcription start site at the 5’ UTR; while the latter two exons, on contig 64, contain the ORF

encoding an allatotropin prepropeptide of 119 amino acid residues and fragments of the 3’ UTR

(Figure 2B). The mature AT-peptide is 13 amino acids long, between residues 45 and 58 and is

identical to the sequence determined by MALDI-TOF mass spectrometry of extracts of R.

prolixus brains (Ons et.al., 2011; Sterkel et.al., 2011). Possible endoproteolytic cleavage sites:

Arg44

, Lys59

Arg60

, flank the mature AT-peptide that ends with a glycine residue; the signal for

carboxyterminal amidation by peptidyl-glycine-α-amidating monooxygenase (Eipper et. al.,

1992). Another possible endoproteolytic cleavage site: Arg93

and Arg94

downstream the mature

peptide has also been identified but the resulting predicted peptide has not been identified in

extracts of the CNS, and so it is not known if the cleavage site is functional. Analysis of

transcript size via a Northern blot yielded a single band of approximately 1.4kb (Figure 2C).

Based on this analysis and in-silico search, it is predicted that approximately 427bp of the

3’UTR is missing, which must also contain the polyadenylation signal (AATAAA).

28

Figure 2. A) Nucleotide sequence of allatotropin (AT) precursor peptide cDNA in R. prolixus.

The entire open reading frame along with the deduced amino acid sequence is shown directly

below the codons. Initial methionine start codon is capitalized, bolded and double underlined

while the stop codon is bolded. Nucleotides preceding the initial start codon and following the

stop codon represent the 5’ and 3’ untranslated regions (UTRs) respectively. Dashed line

represents the 5’ transcription start site, while the possible signal peptide sequence is double

underlined with the possible cleavage site indicated by a bolded asterisk. The boxed region

depicts the mature AT peptide, with the possible dibasic proteolytic cleavage sites shaded in

grey. C terminal glycine residue (Gly58) required for amidation is single underlined. B)

Schematic representation of Rhopr-AT gene organization. The gene is comprised of three

exons- depicted by boxes representing number of nucleotides contained in each; and two

introns- depicted by solid lines. The start (ATG) and stop (TGA) codons are represented by

upward arrows. C) Northern blot of Rhopr-AT transcript hybridised with a complementary

digoxigenin-labelled anti-sense RNA probe 426bp in length. The approximate size of the

transcript was determined using markers shown on the right.

29

30

Rhopr-AT prepropeptide and phylogenetic relationship

The ClustalO alignment represents a high degree of conservation for the 13 residue mature

allatotropin peptide sequence. The Rhopr-AT sequence is approximately 75% identical, with a

shared identity of 10 out of 13 residues, to the lepidopteran species - all of which share an

identical mature allatotropin sequence (Table 1, Figure 3). The mature allatotropin sequence

terminates with the motif TARGF which is also highly conserved among insects; exceptions

being, N. vitripennis, B.impatiens and B. terrestris predicted sequences where the amino acid R

is replaced by Y and T. castaneum where terminal F is replaced by Y. The C-terminal motifs of

the prepropeptides also have a high degree of conservation among species of different taxa

(Figure 3). Species such as R. prolixus, D. pulex and A. aegypti have a single prepropeptide

encoding the mature allatotropin, while lepidopteran species generally have multiple

prepropeptides encoding the mature AT sequence along with Allatotropin-like peptides (Table 1

and 2).

Moreover, lepidopteran species also form a monophyletic group when placed on a phylogenetic

tree. The hypothetical and currently unidentified allatotropin prepropeptide sequences of the

Bombus species form a monophyletic group with the predicted N. vitrepennis allatotropin

sequence, while R. prolixus allatotropin prepropeptide forms a monophyletic group with

Culicidae (Figure 4). The arthropod T. castaneum and mollusc A. california form separate

clades rooting the tree.

RT-PCR

CNS and peripheral tissues were dissected from 10 Vth

instar R. prolixus. RNA was extracted

using Trizol Reagant and cDNA for individual tissues was synthesised. An aliquot of this

single-stranded cDNA was used as a template in a subsequent PCR reaction used to amplify a

553bp segment of Rhopr-AT using gene specific primers. Rp49 was used as the reference gene.

Highest level of expression was observed in the CNS and pool of fat body trachea (Figure 5);

the latter suggesting the presence of peripheral neuroendocrine cells expressing allatotropin. In

only one trial, faint bands were also seen in the immature testes.

31

Figure 3. ClustalO alignment of Rhopr-AT prepropeptide sequence with allatotropin

prepropeptide sequences from other arthropods. Identical amino acid sequences are shaded in

black, while similar amino acids are highlighted in grey. The mature allatotropin sequence is

denoted by a red bar above. The following sequences were used to create the alignment:

Manduca sexta I, Mythimna separata, Mythimna unipuncta, Spodoptera frugiperda I,

Spodoptera exigua, Helicoverpa armigera I, Samia ricini I, Daphnia pulex, Clostera

anastomosis, Aedes aegypti, Anopheles gambiae, Tribolium castaneum, Bombyx mori,

predicted hypothetical protein: Nasonia vitripennis, predicted hypothetical protein: Culex

quinquefasciatus, predicted hypothetical protein: Acyrthosiphon pisum, predicted hypothetical

protein: Bombus impatiens, predicted hypothetical protein: Bombus terrestris.

32

33

Figure 4. Phylogenetic relationship of allatotropin prepropeptide sequence with other insect

sequences. The maximum parsimonius tree was constructed using Close-Neighbor-Interchange

(CNI) analysis where each node represents a percentage corresponding to 1000 bootstrap

replicates. The analysis involved 27 amino acid sequences and was conducted in MEGA5.

34

35

Figure 5. RT-PCR representing the expression profile of the Rhopr-AT transcript in unfed Vth

-

instar R. prolixus. N = 3. Tissues were dissected and the total RNA extracted from 10 insects.1)

CNS, 2) dorsal vessel, 3) foregut, 4) salivary glands, 5) fat body and trachea, 6) anterior midgut,

7) posterior midgut, 8) Malpighian tubules, 9) hindgut, 10) immature testes, 11) immature

ovaries, 12) No template control. A 229bp ribosomal protein 49 (Rp49) was also amplified as a

positive control to test the quality of the cDNA.

36

37

Immunohistochemistry - Phenotypic expression of allatotropin-like immunoreactivity in the

CNS and peripheral tissues

CNS

Intense allatotropin-like immunoreactivity was observed throughout the CNS and some

peripheral tissues of R. prolixus, with the greatest number of intensely-stained cell bodies being

present on the dorsal surface of the brain (Figure 6A, 8A). A cluster of approximately 30-35

cells along the dorsal lateral (dl) regions of each sub-optic lobe (ol) was observed. Varicose

axon terminals were seen in the optic lobes, though the origin of these projections could not be

traced. Moreover, a cluster of 10 bilaterally-paired cells were seen along the dorsal midline (dm)

of the protocerebrum and 3 bilaterally-paired cells were seen along the dorsal anterior midline

(dam) of the protocerebrum (Figure 6A, 8A). The dm cells are likely median neurosecretory

cells. Furthermore, 3 bilaterally-paired dorsal posterior medial cells (dpm) and 2 bilaterally-

paired dorsal posterior lateral cells (dpl) were seen (Figure 6A, 8A). Most of the cells were seen

in the protocerebrum with a few in the tritocerebrum. The ventral surface of the brain had fewer

cells in comparison (Figure 8B). These include clusters of 8 bilaterally-paired cells along the

ventral medial region (vm) in close proximity to 6 bilaterally-paired cells anterior to the

tritocerebrum surrounding the oesophageal canal. Three bilaterally-paired posterior-lateral cells

(pl) were also present in the tritocerebrum. Finally, a cluster of 8 cell bodies at the base of the

optic lobes (ol) were observed in each hemisphere (Figure 8B).

Characteristic bilaterally-paired cell bodies were also observed in each ganglion: the

suboesophageal ganglion (SOG), prothoracic ganglion (PRO) and mesothoracic ganglionic mass

(MTGM). Cells in the SOG were concentrated along the midline; however some cell clusters

were also seen along the lateral surfaces (Figure 6B, 8A). Particularly, 2 distinct pairs of dorsal-

medial (dm) intensely-stained cells (whose projections travelled anteriorly and then laterally

before being lost in the neuropil) were seen dotted by approximately 7 posterior smaller cells at

the dorsal posterior medial region (pm) of the SOG (Figure 6B, 8A). Along the ventral surface 2

brightly-stained cells, surrounded by smaller lightly stained cells were seen anterior to the

neuropil; while 4 pairs of dorso-ventral (dv) bilaterally-paired cells bordered the dorsal medial

intense staining cells (Figure 6B, 8A). Clusters of cells were also seen just posterior to the brain

and a pair of bilaterally-paired cells was seen at the posterior medial region (pm) of the SOG

38

(Figure 8A). Furthermore, varicose axonal processes originating from the posterior SOG

traverse through the PRO and MTGM where they bifurcate and give rise to dense neuropil

regions (Figure 6D, 7A, 8A). These projections are contained within the CNS in the PRO and

SOG as they could not be traced into peripheral nerves, hence suggesting they are interneurons.

In the PRO, 6 anterior-lateral cells (al) and 2 pairs of anterior-medial cells (am) were seen on the

dorsal surface (Figure 6D, 8A), and 2 pairs of anterior-medial cells (am) on the ventral surface

(Figure 8A). Bilaterally-paired ventral medial (vm) and posterior ventral medial (pvm) cells

were also observed in the PRO in close proximity of the axonal projections (Figure 6D, 8A).

The MTGM was more densely populated with cells than the PRO with cell clusters seen around

the axonal processes, which again terminated in dense neuropil regions within the ganglion

(Figure 7A, 8A). Bilaterally-paired cell bodies were observed along the anterior region of the

MTGM. Moreover, a cluster of 8 intensely staining bilaterally-paired cells was seen at the

posterior dorsal surface (pd) of the MTGM and 4 pairs of 3 cells each were also seen at the

posterior of the MTGM (Figure 7A, 7B, 8A).

Periphery

Axonal projections exited the brain and arborized over the retrocerebral complex (CC/CA)

before entering the dorsal vessel (DV) (Figure 6C, 7C). Neurohaemal processes were also

observed on the 2nd

and 3rd

abdominal nerve of the MTGM (Figure 7B, 7E, 8A), as well as on

the trunk nerves (Figure 7D, 8A). The neurohaemal processes observed on the CC/CA, DV

(Figure 7C), abdominal and trunk nerves (Fig. 7B, 7E, 7D) suggest possible neurohormonal

roles for AT in R. prolixus. The accessory duct of the salivary gland showed staining for AT-

like-ir processes across its length, which then proliferated over the principal gland (Figure 9E).

Furthermore, some faint staining was observed over the hindgut (Figure 9A, 9B). However, a

lack of staining in the Malpighian tubules (Fig. 9A, 9C, 9D) challenges the recent report of AT-

like peptide production by the tubules and staining as reported in another triatomine bug,

Triatoma infestans (Santini et. al., 2007, 2009; and see later under Discussion).

39

Figure 6. Confocal images showing AT-like immunostaining in cells of IVth

instar Rhodnius

prolixus CNS, corpus cardiacum and dorsal vessel. A) Brain and anterior suboesophageal

ganglion (SOG). B) SOG. C) Corpus Cardiacum (CC) and dorsal vessel (DV). D) Prothoracic

ganglion (PRO). Arrows point to CC and DV. Scale bars represent 100µm.

40

41

Figure 7. Confocal images showing AT-like immunostaining in cells of IVth

instar Rhodnius

prolixus CNS and peripheral nerves. A) Mesothoracic ganglionic mass (MTGM). B) Abdominal

nerves exiting the MTGM. C) Dorsal vessel (DV). D) Trunk nerve. E) Abdominal nerves. AT-

like immunoreactivity along the CC, DV and nerves is depicted by arrows. N = 60. Scale bars

represent 100µm.

42

43

Figure 8. Camera lucida map depicting distribution of AT-like immunoreactive cells within the

CNS. Open cells are best seen from the ventral surface, while filled cells are best seen from the

dorsal surface. Scale bar represents 200µm. A) Dorsal view. B) Ventral view of the brain. Some

groups have been labelled for clarity in text description. dam = dorsal anterior midline, dm =

dorsal midline, dl = dorsal lateral, ol = optic lobe, dpl = dorsal posterior lateral, dpm= dorsal

posterior medial, dv = dorso-ventral, pm= posterior medial, al = anterior lateral, am = anterior

medial, pd = posterior dorsal, pl = posterior lateral, vm = ventral medial, pvm = posterior

ventral medial.

44

45

Figure 9. Confocal images showing AT-like immunoreactivity in peripheral tissues of IVth

instar

Rhodnius prolixus. A) Posterior midgut (PMG), Malpighian tubules (MT) and hindgut (HG). B)

HG. C) PMG and MT. D) Magnified MT. E) Salivary gland (SG). AT-like immunoreactive

processes are depicted by arrows while areas devoid of staining (notably the MTs) are depicted

by arrowheads. Scale bar represents 100 µm. N= 60.

46

47

Hindgut Assay via Force Transducer

The physiological role of Rhopr-AT upon the hindgut was tested by measuring changes in

longitudinal contractions of the preparation using a force transducer. This tissue was chosen

since there are some faint AT-like immunoreactive processes along its surface. Application of

Rhopr-AT at doses ranging from 10-15

M to 10-6

M resulted in no significant change from the

basal conditions (Figure 10, 11). To test that the bioassay was viable and robust, and the

negative results valid, a positive control was performed. The application of another myoactive

peptide, Rhopr-kinin 2 (10-11

M to 10-6

M), was able to induce forceful contractions of the

hindgut (Figure 10, 11), with maximum effect at 10-8

M Rhopr-kinin 2. Rhopr-AT was also

tested on Vth

instar hindgut to confirm the negative results obtained from IVth

instar hindguts.

Doses of Rhopr-AT also failed to elicit any significant change in contractions. The negative

results obtained were again validated using Rhopr-Kinin 2 (10-11

M to 10-6

M). A sample trace for

both peptides at the same dose is shown in Figure 10.

Heart Impedance Assay

Rhopr-AT (10-12

M to 10-6

M) had no significant effect on heart-beat frequency. No notable

change in frequency was seen after the application of a Rhopr-AT (Figure 12). The myotropic

pentapetide proctolin, however, did yield a large increase in frequency, as seen in Figure 12B.

48

Figure 10. Sample traces depicting effects of different peptide doses on IVth

and Vth

instar

hindguts. Upward arrowhead represents the application of a dose, while the downward arrow

represents saline wash. A and B represent the effect of 10-7

M Rhopr-AT and 10-7

M Rhopr-Kinin

2 upon IVth

instar hindgut respectively. C and D represent the effect of 10-7

M Rhopr-AT and 10-

7M Rhopr-Kinin 2 upon V

th instar hindguts. Large deflections after wash off indicate additional

aggressive washing. N=5.

49

50

Figure 11. Dose-response curves representing the change in longitudinal contractions/tension of

IVth

instar R. prolixus hindgut after application of different doses of Rhopr-AT and Rhopr-kinin

2. N=5.

51

10- 1 6 10- 1 4 10- 1 2 10- 1 0 10- 8 10- 6 10- 40

5

10

15

20

25Rhopr-AT

Rhopr-Kinin 2

Dose (M)

Ten

sio

n (

mg

)

52

Figure 12. A) Dose-response curve for Rhopr-AT activity on heart-beat of R. prolixus. No

significant change in frequency was observed. N=5. B) Sample traces of spontaneous heart-beat

in saline, Rhopr-AT and the myoactive pentapeptide, proctolin. Scale bar represents 30s.

53

A)

B)

54

DISCUSSION

Recent availability of vector genomes and EST databases has facilitated the process of

identification and isolation of genes that encode neuropeptides, neurohormones and

neuromodulators within species. Using an in-silico approach the allatotropin gene for the

triatomine bug, Rhodnius prolixus, was identified and subsequently cloned to reveal a cDNA

fragment of 973bp encoding one mature amidated allatotropin peptide. The amidated

tridecapeptide is identical to the Rhopr-AT sequence determined by MALDI-TOF mass

spectrometry of extracts of R. prolixus brains (Ons et.al., 2011; Ons et.al., 2009). A Northern

blot further certified the presence of a single mRNA encoding a single mature allatotropin

peptide, 1.4kb in length. Absence of multiple bands in the Northern blot, along with single hits

from the BLAST results also confirms that Rhopr-AT is a single transcript encoded by one gene

without splice variants.

The expression profile for Rhopr-AT via RT-PCR revealed high transcript levels in the CNS and

pool of fat body and trachea alluding to the presence of possible peripheral neuroendocrine cells

that also express allatotropin. Furthermore, immunohistochemical analysis of R. prolixus tissues

displayed AT-like-ir cells well-dispersed throughout the CNS along with AT-like-ir processes

projecting to peripheral tissues. Faint staining was observed over the posterior midgut and

hindgut, while no staining was observed in the Malpighian tubules. Allatotropin-like

immunoreactive axons in the salivary nerve traverse along the accessory duct before forming a

nerve plexus over the principal gland of the salivary gland.

Contraction assays with the force transducer, to determine the effect of Rhopr-AT upon the

hindgut yielded negative results for both IVth

and Vth

instars. To verify that the negative results

obtained were due to inactivity of Rhopr-AT and no other artefact, a positive control using

Rhopr-kinin 2 was performed. The myotropic peptide, Rhopr-kinin 2 was able to induce robust

longitudinal contractions in the hindguts of both IVth

and Vth

instars. Considering the elaborate

nerve plexus over the dorsal vessel, it was postulated that Rhopr-AT might stimulate

contractions of the heart. Interestingly though, no such effect was observed. The negative results

yielded from the heart impedance assay were validated with the myotropic pentapeptide

proctolin, which was able to dramatically increase the frequency of contractions. The

55

allatotropic activity of Rhopr-AT on JH production cannot be tested at this time since the R.

prolixus JH is yet to be chemically identified.

Allatotropins (ATs) first isolated from the heads of adult Manduca sexta (Kataoka et al., 1989)

have since been isolated and characterised extensively in several other lepidopteran species. The

AT gene is transcribed as multiple mRNA isoforms within most lepidopterans; each isoform

differing from another as a consequence of alternative splicing and encoding the mature

amidated tridecapeptide AT along with allatotropin-like peptides (ATL) (Abdel-latief et al.,

2003; Nagata et al., 2012; Sheng et al., 2007; Taylor et al., 1996; Truesdell et al., 2000; Yin et

al., 2005). The phenomenon of multiple AT mRNA isoforms is limited to the lepidopteran order

and characterization and prediction of AT in various oth