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The Role of Octopamine and Tyramine in the Adult
Female Reproductive System of Rhodnius prolixus
by
Sam Hana
A thesis submitted in conformity with the requirements
for the degree of Master of Science
Cell and Systems Biology
University of Toronto
© Copyright by Sam Hana 2017
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The Role of Octopamine and Tyramine in the Adult
Female Reproductive System of Rhodnius prolixus
Sam Hana
Master of Science
Cell and Systems Biology
University of Toronto
2017
Abstract
Octopamine and tyramine are neuroactive chemicals involved in
many physiological
processes acting as neurotransmitters, neuromodulators and
neurohormones. Octopamine and
tyramine modulate reproduction in insects. In Rhodnius prolixus,
octopamine decreased the
amplitude and reduced the RhoprFIRFa-induced oviduct contraction
in a dose-dependent manner,
whereas tyramine only reduced the RhoprFIRFa-induced
contractions. Also, octopamine and
tyramine reduced the frequency and abolished bursal contractions
at higher concentrations.
Octopamine also increased the levels of cAMP in the oviducts, an
effect blocked by phentolamine.
Dibutyryl cAMP mimicked the effects of octopamine at the bursa,
suggesting that octopamine may
act by an Octβ-receptor, a known GPCR. The cDNA sequences of
RhoprOctβ2-R and RhoprTyr1-
R have been cloned and characterized; the receptor transcripts
are expressed in all female
reproductive tissues. Injection of octopamine and tyramine into
mated and fed adult females
increased oogenesis. Overall, octopamine and tyramine modulate
the female reproductive tissues
leading to successful laying of eggs.
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Acknowledgments
I would like to first offer my deepest thanks to my mentor, Dr.
Angela Lange. Her excellent
guidance and supervision were instrumental in my success. She
motivated and inspired me to do
my best throughout this journey. I am fortunate to graduate
under your supervision knowing that
I was taught from the best.
I want to acknowledge and thank my committee, Dr. Ian Orchard
and Dr. Adriano Senatore for
their input in my research. Dr. Senatore, thank you very much
for your helpful tips and feedback
early on in my degree. Dr. Ian Orchard, I would like to thank
you for reading, editing, and
providing feedback on the research articles and this thesis.
To all members of the Lange lab, I want to thank you for being
part of this journey. I especially
want to acknowledge those that taught, guided and supported me
in my research. You have been
great colleagues and I am glad to have known you all. We have
created wonderful memories.
And lastly, there was no limit to the amount of love and support
my family has given me. Thank
you father and mother for your sacrifices every day, I am
blessed to have you. Thank you to my
brothers for the love and support throughout this journey. Thank
you God for the countless
blessings.
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Table of Contents
Abstract
..........................................................................................................................................
ii
Acknowledgments
.........................................................................................................................
ii
Organization of the Thesis
.........................................................................................................
vii
List of Figures and Supplementary Tables
..............................................................................
viii
List of Abbreviation
.......................................................................................................................x
Chapter 1: General Introduction
.................................................................................................1
Neuroactive chemicals
.....................................................................................................................1
Biogenic amines
...............................................................................................................................1
Octopamine
......................................................................................................................................2
Tyramine
..........................................................................................................................................3
Rhodnius prolixus: a vector of Chagas disease
................................................................................7
The female reproductive system
......................................................................................................8
Anatomy
...........................................................................................................................................8
Central nervous system innervation to the reproductive system
...................................................12
Reproductive processes
..................................................................................................................12
Oogenesis
...........................................................................................................................12
Ovulation
............................................................................................................................13
Fertilization and oviposition
..............................................................................................13
Neuroactive chemicals control reproduction in females
................................................................14
Significance....................................................................................................................................15
Thesis objective
.............................................................................................................................16
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References
......................................................................................................................................18
Chapter 2: Octopamine and tyramine regulate the activity of
reproductive visceral
muscles in the adult female blood-feeding bug, Rhodnius
prolixus. ...................................23
Abstract
..........................................................................................................................................24
Introduction
....................................................................................................................................25
Materials and Methods
...................................................................................................................26
Results
............................................................................................................................................29
Discussion
......................................................................................................................................44
References
......................................................................................................................................47
Chapter 3: Cloning and functional characterization of
Octβ2-Receptor and Tyr1-
Receptor in the Chagas disease vector, Rhodnius prolixus.
.................................................50
Abstract
..........................................................................................................................................51
Introduction
....................................................................................................................................52
Materials and Methods
...................................................................................................................56
Results
............................................................................................................................................59
Discussion
......................................................................................................................................79
References
......................................................................................................................................84
Supplementary Material
.................................................................................................................90
Chapter 4: Octopamine and tyramine induce egg-laying in Rhodnius
prolixus. ...................93
Abstract
..........................................................................................................................................94
Introduction
....................................................................................................................................95
Materials and Methods
...................................................................................................................97
Results
..........................................................................................................................................101
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Discussion
....................................................................................................................................106
References
....................................................................................................................................108
Chapter 5: General Discussion
.................................................................................................111
The role of octopamine and tyramine in the reproductive system
...............................................111
Reproductive visceral muscle
......................................................................................................112
▪ Oviducts
...........................................................................................................................112
▪ Bursa
................................................................................................................................114
Reproductive processes
................................................................................................................117
▪ Direct
................................................................................................................................117
▪ Indirect
.............................................................................................................................117
Summary
......................................................................................................................................121
Future directions
..........................................................................................................................122
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Organization of the Thesis
The thesis is broken into 5 chapters. Chapter 1 provides a
general introduction for the thesis.
Chapter 2 is organized as a research journal article and it
focuses on octopamine and tyramine
regulation of reproductive visceral muscles. Chapter 2 has been
published in the Journal of
Experimental Biology (Hana and Lange, 2017). Chapter 3 focuses
on RhoprOctβ2-R and
RhoprTyr1-R cDNA receptor cloning, functional characterization
and expression in the
reproductive system. Chapter 3 is also organized as a research
article that has been submitted
for publication in Frontiers in Physiology. Chapter 4 is a short
report on the effects of injected
octopamine and tyramine on egg-laying. Chapter 5 is for general
discussion and connects all the
chapters.
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List of Figures and Supplementary Tables
Chapter 1: General Introduction
Figure 1. The biosynthetic pathway of octopamine and tyramine
..................................................4
Figure 2. The anatomical structures of the adult female
reproductive system .............................10
Chapter 2: Octopamine and tyramine regulate the activity of
reproductive visceral
muscles in the adult female blood-feeding bug, Rhodnius
prolixus
Figure 1. Octopamine and tyramine on rhythmic contractions of
the oviducts............................32
Figure 2. Octopamine and tyramine on RhoprFIRFa-induced
contractions……………….........34
Figure 3. Octopamine and tyramine on rhythmic contractions of
the bursa……………….........36
Figure 4. Phentolamine inhibits octopamine in the lateral
oviducts…………………..…...........38
Figure 5. Phentolamine blocks octopamine in the
bursa…………………………..………........40
Figure 6. Yohimbine fails to block tyramine in the
bursa…………………………..……..........42
Chapter 3: Cloning and functional characterization of
Octβ2-Receptor and Tyr1-Receptor
in the Chagas disease vector, Rhodnius prolixus.
Figure 1. Classification of octopamine and tyramine
receptors....................................................56
Figure 2. RhoprOctβ2-R cDNA sequence
………………………………..………..……...........64
Figure 3. RhoprTyr1-R cDNA sequence
……………………………………………..…...........66
Figure 4. Phylogenetic tree of insect octopamine and tyramine
receptors …………...…….......68
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Figure 5. Multiple sequence alignment of insect Octβ2-Rs
…………………………..…….......70
Figure 6. Multiple sequence alignment of insect Tyr1-Rs
…………………………...……........72
Figure 7. Functional characterization of R. prolixus Octβ2
receptor……………………..……..74
Figure 8. Functional characterization of R. prolixus Tyr1
receptor………………………..……76
Figure 9. Spatial expression of RhoprOctβ2-R and RhoprTyr1-R.
……………………...……..78
Table S1. Primers for the amplification of the receptor
fragments ……………….....…..…..….91
Table S2. Primers for the amplification of 3’ region of the
receptors…………………….....….91
Table S3. Primers for the amplification of 5’ region of the
receptors ……………...……...…...92
Table S4. Primers used for the mammalian expression vector
preparation ……………....……92
Table S5. Primers used for RT-qPCR analysis of receptor
transcripts………………....….……93
Chapter 4: Octopamine and tyramine induce egg-laying in Rhodnius
prolixus
Figure 1. The protocol for the egg-laying
assay……………………………………………….101
Figure 2. The result of octopamine injection on the number of
eggslaid…………….………..103
Figure 3. The effect of tyramine injection on the number of eggs
laid………………...………105
Chapter 5: General Discussion
Figure 1. Model showing the effects of octopamine on the
oviducts and the bursa…………...116
Figure 2. Model for the possible effects of octopamine and
tyramine on egg production…….120
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List of Abbreviation
ANOVA analysis of variance
AST Allatostatin
BLAST basic local alignment search tool
bp base pairs
CA corpora allatum
Ca2+ calcium
cAMP 3’-5’-cyclic adenosine monophosphate
CC corpus cardiacum
cDNA complementary deoxyribonucleic acid
CNS central nervous system
EC50 half maximal activation concentration
FLPs FMRFamide-like peptides
GDP guanosine diphosphate
GPCR G-protein coupled receptor
GTP guanosine triphosphate
HEK293/CNG human embryonic kidney cells expressing cyclic
nucleotide-gated ion
channel
IP3 inositol triphosphate
JH juvenile hormone
mNSCs median neurosecretory cells
mRNA messenger ribonucleic acid
MS Myosuppressin
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MTGM mesothoracic ganglionic mass
NPF Long neuropeptide F
ORF open reading frame
PBS phosphate-buffered saline
PRO prothoracic ganglion
RT-qPCR quantitative reverse transcription Polymerase Chain
Reaction
RACE rapid amplification of cDNA ends
SEM standard error of the mean
SOG suboesophageal ganglion
TM transmembrane domain
UTR
untranslated region
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Chapter 1: General Introduction
Neuroactive chemicals
Insects display a wide range of behaviours important for
maintaining survival and
ultimately reproductive success. Insects utilize the nervous
system, composed of the brain and the
ventral nerve cord, to monitor external environmental cues and
coordinate internal processes. This
complex coordination is enabled by a large network of
functionally diverse neurons and supporting
cells. Different neuroactive chemicals are utilized by the
nervous system to transmit signals from
neuron to neuron and neuron to target tissues, thereby allowing
the flow of information within the
organism. These neuroactive chemicals are classified as
neurotransmitters, neurohormones and
neuromodulators (Klowden, 2013). Neurotransmitters are
specifically released into the synaptic
cleft causing changes in the post-synaptic membrane potential.
Glutamate is known as the classic
excitatory neurotransmitter at neuromuscular junctions in
insects (Jan and Jan, 1976). The effect(s)
of neurotransmitters are transient due to re-uptake, enzymatic
degradation and diffusion in the
synaptic gap. Neurohormones, synthesized by neurosecretory
cells, are released into the
hemolymph and function as circulating hormones. Neurohormones
modulate many peripheral
tissues and have longer-lasting effects. For example,
adipokinetic hormone released from the
corpus cardiacum regulates energy levels in insects (Orchard and
Lange, 1983). Lastly,
neuromodulators, released by neurons, modify the transmission in
other synapses and/or modify
activity at target tissues. Neuromodulators can affect the
excitability of post-synaptic membranes
and modulate the release of neurotransmitters from the
pre-synaptic neurons. For example, at high
concentrations, serotonin can enhance the excitatory responses
evoked by electrical stimulation of
the antennal nerves, whereas at low concentrations, serotonin
reduces theses responses in Manduca
sexta (Kloppenburg and Hildebrand, 1995).
Biogenic amines
Biogenic amines are a class of organic neuroactive chemicals,
mostly obtained from amino
acids and characterized by having low molecular weights and
amine functional group(s). In insects,
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octopamine, tyramine, serotonin (5-HT), dopamine and histamine
are examples of the most
common and widely studied biogenic amines. Biogenic amines can
be utilized by the nervous
system for the transmission of quick, private and short-lasting
messages (Orchard et al., 2001).
Biogenic amines are mainly distributed in the central nervous
system (CNS), but are also found in
projections to peripheral tissues suggesting their diverse
physiological roles in insects (Blenau and
Thamm, 2011; Monastirioti, 1999; Nässel, 1999; Nässel and
Elekes, 1992; Roeder, 2005). They
can also act as neurohormones or neuromodulators. Serotonin is
released into the hemolymph
during feeding in Rhodnius prolixus to initiate diuresis by
stimulating rapid tubule secretion
(Maddrell et al., 1991). In Periplaneta americana, dopamine
stimulates the release of the fluid
component of saliva and serotonin stimulates the release of the
proteinaceous components of saliva
(Just and Walz, 1996). Histamine, in Musca domestica and
Calliphora erythrocephala, is a
neurotransmitter involved in olfaction as its been shown to be
released from photoreceptor cells
(Hardie, 1987). These are just some examples of the
physiological effects of these biogenic amines.
Octopamine
Octopamine was discovered in the salivary gland of the octopus,
Octopus vulgaris
(Erspamer, 1948). Octopamine is a versatile neuroactive chemical
derived from the hydroxylation
of tyramine by tyramine β-hydroxylase (Fig. 1). There is an
immense amount of data that confirms
that octopamine acts as a neurotransmitter, neuromodulator and a
neurohormone in insects, hence
the its versatility (Orchard, 1982). Octopamine is structurally
and functionally similar to
noradrenaline of vertebrates; octopamine is the “adrenergic” of
insects (Roeder, 1999; Roeder,
2005). In the CNS, octopamine’s content is 3-7 times higher than
tyramine (Lange, 2009). In the
adult D. melanogaster, octopamine is found throughout the CNS
with processes innervating
regions such as the ventral nerve cord, subesophageal ganglia,
protocerebrum, central complex,
mushroom bodies and the optic lobe (Busch et al., 2009).
Multiple groups of neurons containing
octopamine have been characterized, like the ventral median
paired (VPM), ventral unpaired
median (VUM) and dorsal unpaired median neurons (DUM) (Busch et
al., 2009). In Schistocerca
gregaria and P. americana, octopamine is found in the ganglia of
the ventral nerve cord and is
also found in the optic lopes (Evans, 1978). In both species,
octopamine was also found to be
associated with the corpora cardiaca (Evans, 1978). Octopamine
is involved in a plethora of
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physiological processes thereby influencing communication,
signaling and behaviour in insects.
To briefly state a few roles, octopamine is important in
aggression, reproduction, learning and
memory (Farooqui, 2012; Ohta and Ozoe, 2014; Roeder, 1999;
Roeder, 2005).
Tyramine
Tyramine was not originally thought be being a neurotransmitter
but merely thought to be
the metabolic precursor of octopamine; however, tyramine has
been established as an independent
bioactive chemical in insects synthesized via the
decarboxylation of the amino acid tyrosine
(Lange, 2009) (Fig. 1). In Drosophila melanogaster, tyrosine
decarboxylase 2 is responsible for
synthesizing tyramine in neuronal cells, whereas tyrosine
decarboxylase 1 synthesizes non-
neuronal tyramine in peripheral tissues (Cole et al., 2005). The
distribution of tyramine specific
neurons, those that do not also contain octopamine, has been
described in insects. In S. gregaria,
immunoreactive tyramine neurons are found in the medulla, the
protocerebral bridge, the antennal
lobes, subesophageal ganglion and the neuropile (Homberg et al.,
2013; Kononenko et al., 2009).
In D. melanogaster larva, tyramine-specific neurons are found in
the brain, thoracic ganglia and
abdominal ganglia (Monastirioti et al., 1995).
Tyramine-containing neurons that also contain
octopamine, have been found in the suboesophageal ganglia and
thoracico-abdominal ganglia
(Nagaya et al., 2002). These neurons are characterized as
ventral unpaired medial neurons (VUM)
and the tyramine specific pairs of dorsal lateral neurons in the
abdominal ganglia (Nagaya et al.,
2002). VUM and DUM neurons, which also contain octopamine,
innervate skeletal muscles and
other peripheral target tissues (Lange, 2009). Physiologically,
tyramine along with its metabolic
successor, octopamine, have been shown to have similar effects
in invertebrates (Roeder, 1999;
Roeder, 2005); however, tyramine has been reported to exhibit
its own physiological actions
independent of octopamine. This was confirmed by the discovery
of tyramine specific receptors.
For example, elevation of tyramine attenuates locomotion in D.
melanogaster larva lacking
octopamine and octopamine feeding rescues locomotion in mutant
flies (Saraswati et al., 2004).
Tyramine has been shown to be involved in the regulation of many
physiological processes, such
as reproduction, aggression, feeding and locomotion (see Ohta
and Ozoe, 2014).
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Figure 1. The biosynthetic pathway for the synthesis of
octopamine and tyramine. Tyramine is
produced by the decarboxylation of tyrosine, while octopamine is
produced by the hydroxylation
of octopamine. Figure obtained from (Lange, 2009)
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G-protein-coupled receptors (GPCRs)
G-protein-coupled receptors (GPCRs), also known as
seven-transmembrane domain
receptors (7TMs), are a large family of receptor proteins that
bind ligands extracellularly leading
to receptor activation and thereby eliciting intracellular
signal transduction (Kristiansen, 2004).
GPCRs are coupled to specific G-proteins which are an integral
component in initiating signal
transduction pathways. GPCR ligands include biogenic amines,
neuropeptides, glycoproteins,
photons, taste molecules, odorants, hormones + pheromones and
odorants (Kristiansen, 2004). All
GPCRs are characterized by having an extracellular N-terminus,
7TMs creating three extracellular
loops along with three intracellular loops and an intracellular
C-terminus. The N-terminus is
glycosylated, and this is critical for cell surface expression
(Kristiansen, 2004). The 7TM domains,
7 α-helices, span the plasma membrane usually creating a pocket
in which ligands interact with
the side chain amino acids (Kristiansen, 2004). The
extracellular loops can also participate in
ligand interaction and/or binding (Kristiansen, 2004). The
cytosolic loops contain serine and
threonine residues which are potential phosphorylation sites by
protein kinases. Protein kinases
serve as a method of GPCR desensitization (Kristiansen, 2004).
The C-terminus contains cysteine
residues which serve as a site for palmitoylation.
Palmitoylation anchors the C-terminus to the
cytosolic plasma membrane. Palmitoylation allows for normal
processing of the receptor and
accessibility of the C-terminus to kinases and regulatory
proteins (Kristiansen, 2004). G-proteins
are divided into two main classes, heterotrimeric G-proteins and
small cytoplasmic G-proteins.
Heterotrimeric G-protein complexes are composed of α, β and γ
subunits. G-proteins are closely
associated with GPCRs at the amphiphatic α-helices of TM5 and
TM6 (Kristiansen, 2004). They
are also believed to interact with the intracellular loops (ICL2
and ICL3) and the C-terminus
(Kristiansen, 2004).
The binding of a ligand to the GPCR causes receptor activation
and conformational change
leading to an increased affinity for the G-protein (Kristiansen,
2004). This causes the release of
guanosine diphosphate (GDP) from the α subunit and the binding
of guanosine triphosphate (GTP)
(Kristiansen, 2004). The α subunit, bound to GTP, dissociates
from the βγ complex and leads to
the activation of target proteins, initiating signaling cascades
(Kristiansen, 2004). The activated
GPCR lasts until GTP in α subunit is hydrolyzed to GDP and the
heterotrimeric complex reforms
(Kristiansen, 2004). There are many classes of Gα subunits.
Different subunit lead to different
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activation or inhibition of downstream effectors (Ellis, 2004).
Gαs subunit is known to activate
adenylate cyclase and lead to cAMP elevation, whereas, Gαi/Gαo
inhibits adenylate cyclase (Ellis,
2004).
There are six different subfamilies of GPCRs. Biogenic amines
belong to the rhodopsin-
like (Class A) subfamily of GPCRs. It is the largest family of
GPCRs and members of this family
are characterized by having DRY motif at the end of TM3, and
NPxxY domain in TM7 (Rovati
et al., 2007; White et al., 2012). These residues are important
for protein stabilization and/or G-
protein activation. Biogenic amines noncovalently bind to the
upper part of the 7TM domains. The
binding of these small neuroactive chemicals lies deep between
TM3, TM4, TM5, TM6 and TM7
(Kristiansen, 2004). Many insect biogenic amines receptors have
been characterized and
functionally analyzed. A focus on octopamine and tyramine
receptors is presented in Chapter 3.
The function of these receptors is integral in all animals.
Therefore, these GPCRs are potential
targets of many insecticides.
Rhodnius prolixus: a vector of Chagas disease
Rhodnius prolixus, generally known as one of many kissing bugs,
is a blood-feeding
hemipteran mainly found in South and Central America (Bern et
al., 2011). R. prolixus undergoes
incomplete metamorphosis developing through five nymphal instars
and finally molting into a
sexually capable adult stage (Nunes-da-fonseca et al., 2017). A
blood meal is required for every
incomplete metamorphosis; those that do not obtain a blood meal
attenuate development and
remain in the nymphal stage. R. prolixus typically feed on
mammalian, bird, marsupial and
reptilian blood (Davey, 2007). There are two known populations
of R. prolixus, sylvan and
domestic (Davey, 2007). Sylvan populations occur among the
leaves of palm trees, pteridophytes,
and their hosts’ burrows and nests (Davey, 2007). Domestic R.
prolixus are in close association
with human habitats and living spaces. They are found in
crevices, thatched roofs and small damp
dark spaces (Davey, 2007). R. prolixus are resilient insects,
and nymphal instar stages have been
observed to survive for several months without a blood meal in
colonies grown and maintained in
the laboratory (WHO, 2002).
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R. prolixus are nocturnal, they sense CO2 levels and are
attracted to the mouth and eye
regions of the human host (CDC, 2013). Feeding behaviour is
initiated with the insertion of the
proboscis into the host. A cocktail of chemicals are injected
into the host via the proboscis to
prevent detection, blood coagulation and vasoconstriction (WHO,
2002). Fifth-instar R. prolixus
have been observed to feed for 20 minutes thereby consuming 10
times their unfed body mass
(Orchard, 2006). Taking such a massive blood meal causes a major
physiological disturbance in
R. prolixus. While feeding, diuresis is initiated for the
expulsion of excess water and salts to
compensate for the large blood meal (Orchard, 2006). Trypanosoma
cruzi, a protozoan carried in
the gut of Rhodnius prolixus, is also expelled in the process of
diuresis (Garcia et al., 2007). T.
cruzi is introduced into the host by mucosal membrane or through
the blood stream by scratching
of the punctured area (WHO, 2002).
T. cruzi is known to cause Chagas disease which infects 6 to 7
million people worldwide
(WHO, 2017). Most cases of the disease have been reported in
Latin America (WHO, 2017). In
the United States, the Center for Disease Control and Prevention
estimates that 300,000 people are
carriers of the parasite; these individuals include immigrants
from Chagas disease native regions
(CDC, 2013). In the acute phase of the disease, the parasite is
found in the circulating blood and
fever and swelling at the site of infection are reported. In the
chronic phase, 20 to 30% of infected
individuals can develop life-threatening conditions due to
cardiac and digestive problems
(Kirchhoff and Pearson, 2007). No vaccines have been developed
yet, however there are
treatments for the acute phase of the disease that can eliminate
T. cruzi from the blood of the host
(CDC, 2013).
The female reproductive system
Anatomy
The adult female reproductive system is composed of two ovaries,
two lateral oviducts, a
common oviduct, spermatheca, bursa and the cement gland
(Wigglesworth, 1972) (Fig. 2). The
terminal filament attaches to the body wall. Each ovary contains
seven meroistic telotrophic type
ovarioles connected to a terminal filament. The ovarioles within
the ovary are held together by a
muscular layer known as the peritoneal sheath (Sedra and Lange,
2014). The ovary is the site of
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egg growth and development, oogenesis. Mature eggs in the ovary
are deposited into the oviducts
by a process known as ovulation. Eggs are then carried to the
common oviduct by oviduct
peristaltic contractions. Sperm stored in the spermatheca,
released by spermathecal contractions,
fertilizes the eggs at the common oviduct (Davey, 1958). The
fertilized eggs move to the bursa for
oviposition. The eggs are first coated with secretions from the
cement gland for attachment to
substrates and then laid by strong bursal phasic contractions
(Lococo and Huebner, 1980).
All muscle in insects are striated. Insect muscles can be
divided into skeletal muscle,
visceral muscle, and cardiac muscles (Wigglesworth, 1972).
Skeletal muscles are attached to the
cuticle and serve for locomotion and moulting. Visceral muscles
have only one or commonly no
attachments to the cuticle. Visceral muscles serve to move the
visceral organs, one of which is the
reproductive system. Visceral muscles in the insect are
myogenic, they commonly have slow
rhythmic contractions (Orchard and Lange, 1986). Contractions
can also be neurogenic, these
contraction are initiated by the nervous system enable fine
control of visceral processes (Orchard
and Lange, 1986).
The reproductive system of the adult female R. prolixus is made
of myogenic visceral
muscle and has been well described by Sedra and Lange 2014. The
terminal filament is made up
of a thick muscular structure. The ovary itself is encircled
with a network of muscle fibers. Each
ovariole is surrounded with a criss-cross network of muscle
fibers. The lateral oviducts are
surrounded with first a layer of longitudinal and then circular
muscle fibers. Thicker circular
muscle fibers also found at the common oviduct and the
spermatheca. Thick longitudinal muscles
arranged in a chevron make up the bursa. The cement gland is
largely non-muscular except at the
proximal end of the gland.
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Figure 2. The anatomical structures of the adult female
reproductive system of R. prolixus.
Illustration created by Paul Hong.
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Central nervous system innervation to the reproductive
system
The CNS of R. prolixus is made of a dorsal brain connected to a
ventral suboesophageal
ganglion (SOG) located in the narrow head region (Tsang and
Orchard, 1991). The SOG is
connected to the prothoracic ganglion (PRO) which is then
connected to the mesothoracic
ganglionic mass (MTGM) (Tsang and Orchard, 1991). The MTGM is a
large ganglion because it
houses the fused mesothoracic ganglion, metathoracic ganglion
and abdominal ganglion. The
MTGM sends nerve fibers that innervate regions in the insect’s
abdomen (Insausti, 1994). The
reproductive system is innervated by the trunk nerve (Chiang and
O’Donnell, 2009; Insausti, 1994;
Sedra and Lange, 2014). Various neuroactive chemicals are known
to innervate the reproductive
system of R. prolixus. Immunoreactivity to FMRFamide-like
peptides was detected in process the
ovaries, oviducts, spermatheca and the bursa. In D.
melanogaster, octopamine and tyramine
innervate the peritoneal sheaths in the ovary, oviducts,
spermatheca and the uterus (Middleton et
al., 2006; Rodriguez-Valentin et al., 2006).
Reproductive processes
Oogenesis
As previously stated, hemipterans have meriostic teleotrophic
type ovarioles (Huebner and
Anderson, 1972). Meriostic type ovarioles are characterized by
having nurse cells, nutritive cells,
and germ cells that contribute to the nourishment of the
developing oocyte (Huebner and
Anderson, 1972). Telotrophic type, a subgroup of meriostic
ovarioles, are characterized by having
nurse cells restricted to the apex of the ovariole and provide
nutrients to the developing oocyte by
a nutritive cord (Wigglesworth, 1972). These ovarioles are
subdivided into four main regions: the
terminal filament, the germarium, the vitellarium and the
ovariole stalk (Nunes-da-fonseca et al.,
2017). The germarium region is located below the terminal fiber
that attaches the ovariole. The
germarium is composed of undifferentiated oogonia (germline
cells) and nurse cells
(Wigglesworth, 1972). The oogonia differentiate into an oocytes
and nurse cells. The developing
oocyte, nourished by nurse cells via the nutritive cord and the
surrounding follicular cells, moves
down the ovariole into the vitellarium (Bonhag, 1955). The
vitellarium is where vitellogenesis
occurs. Vitellogenin, egg yolk, is made in the fat body and is
deposited into the hemolymph
(Davey, 1981). Vitellogenin in the hemolymph is transferred to
the oocyte via gaps between the
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13
follicular cells. Vitellogenin is also synthesized in the
follicular cells and directly deposited into
the oocyte (Davey, 1981). Nurse cells also provide vitellogenin
via the nutritive cord. After the
developing egg enlarges, the follicular gaps close and
vitellogenin uptake ends (Patchin and
Davey, 1968). The follicular cells secrete an elastic vitelline
membrane and then form the chorion
(Patchin and Davey, 1968).
Ovulation
After the formation of the chorion, the follicular cells that
surround the mature egg
degenerate. This leaves the egg free to move and in direct
contact with the peritoneal sheath.
Various hormones circulating in the hemolymph and neurochemicals
such as myotropins secreted
by the CNS stimulate contractions in the ovary and oviducts
(Davey, 1967). Contractions in the
ovary are synchronized with relaxation of the lateral oviducts
(Hana and Lange, 2017; Middleton
et al., 2006). This coordination between the ovary and the
lateral oviducts allows the movement of
eggs from the ovary to the lateral oviducts. Peristaltic
contractions aided with lumen secretions in
the lateral oviducts allows the movement of ovulated eggs into
the common oviduct (Masetti et
al., 1994; Sun and Spradling, 2013). Once the eggs are ovulated,
they are immediately fertilized
and laid (Kriger and Davey, 1982).
Fertilization and oviposition
Before the process of fertilization of eggs, spermatozoa must be
first taken up and stored
in the spermatheca (Davey, 1958). The spermatophore, containing
spermatozoa, is deposited into
the bursa of the female during copulation (Davey, 1958). Male
accessory gland secretions leads to
contractions in the common oviduct leading to the movement of
the spermatozoa to the
spermatheca (Davey, 1958). The spermatozoa does not play an
active role in the migration (Davey,
1958). When the egg arrives at the common oviduct, contractions
in the spermatheca cause the
release of spermatozoa onto the chorion coated egg. The
spermatozoa fertilizes the egg via narrow
passages within the chorion known as micropyles (Okasha et al.,
1970). The fertilized egg moves
to the bursa via contractions in the common oviduct. Secretions
from the cement gland coats the
egg and the egg is then deposited onto a substrate by strong
phasic bursal contractions (Lococo
and Huebner, 1980).
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14
Neuroactive chemicals control reproduction in females
Rhodnius prolixus, like all blood-feeding insects, require blood
meals for reproductive
maturity. Despite having a fully functioning reproductive
system, female adults must generally be
fed in order to mate (Buxton, 1930). Once they are fed, they can
synthesize and lay viable eggs.
Mating introduces male factors that accelerate reproductive
processes in females (Davey, 1965).
At the germarium, an oocyte’s growth is directly regulated by
its genome and the content
of its species-specific information is known as euplasm; the
euplasm is synthesized in the nurse
cells (Chapman, 2013). At the vitellarium, yolk uptake supresses
DNA transcription and nuclear
processes. Juvenile hormone (JH) secreted from the corpus
allatum acts on the fat body to
synthesize vitellogenin and leads to the formation of gaps
between follicular cells (patency)
(Davey, 2007; Davey et al., 1974). These follicular gaps are
needed for vitellogenin uptake by the
oocyte (Davey et al., 1974). The synthesis and release of JH is
under the control of allatotropins
and allatostatins in the brain (Davey, 1987; Teal, 2002). The
brain, acting via neurosecretory cells,
can signal for JH release when mating and feeding have occurred
(Davey, 2007). Antigonadotropin
released from neurosecretory cells in the abdomen counteract the
effects of JH on follicle cells and
prevent the formation of gaps between follicular cells (decrease
patency) (Davey and Kuster,
1981). Recently, it was shown that injected RhoprShortNPF
(NNRSPQLRLRFamide),
RhoprFMRFa (GNDNFMRFamide) and RhoprFIRFa (AKDNFIRFamide)
increases the number
of eggs produced, i.e. increases oogenesis (Sedra and Lange,
2016). In contrast, injections of
RhoprMS (pQDIDHVFMRFamide) and RhoprAST-2 (LPVYNFGLamide)
reduced egg
production (Sedra and Lange, 2016). Therefore, FMRFamide-like
peptides (FLPs) are important
for controlling the rate of oogenesis. The mechanisms where this
occurs and not known.
Ecdysteroids, produced from the ovary, do not affect
vitellogenin content in the hemolymph and
do not increase egg production (Davey, 2007).
At the end of vitellogenesis, ecdysteroids synthesized from the
follicular cells are thought
to be essential for ovulation (Chapman, 2013). Circulating
ecdysteroids elicit the release of a
myotropic ovulation hormone from the corpus cardiacum of the
brain (Ruegg et al., 1981). A
myotropin was found to be synthesized from ten neurosecretory
cells in the pars intercerebralis
(Ruegg et al., 1981). This myotropin leads to an increase in
muscle contractions in the ovary and
thereby increases the rate of ovulation (Davey, 1967; Ruegg et
al., 1981). A FLP has been
suggested to the ten neurosecretory cells (Sevala et al., 1992).
The peak of FLP release, five days
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15
post-feeding, parallels the peak of myotropin release in the
gonadotrophic cycle (Sevala et al.,
1992). Therefore, the myotropin is thought to be related to FLPs
(Sevala et al., 1992). Furthermore,
other neuropeptides and biogenic amines can modulate ovulation
by controlling lateral oviduct
contractions. For example RhoprFIRFa and RhoprFMRFa have been
shown to increase ovariole
and lateral oviduct contractions (Sedra and Lange, 2014). The
myoinhibitors, RhoprAST-2 and
RhoprMIP-4, inhibited oviduct contractions (Sedra et al., 2015).
Octopamine’s role in oviduct
relaxation has been shown in other insects. In D. melanogaster,
octopamine stimulated peritoneal
sheath contractions and relaxed the oviducts (Middleton et al.,
2006; Rodriguez-Valentin et al.,
2006). In Locusta migratoria, serotonin, acting as a
neuromodulator, elicits an increase in
amplitude of oviduct contractions (Lange, 2004). After
ovulation, the eggs are moved through the
oviducts by peristaltic and phasic contractions. For
fertilization to occur, the spermatheca must
contract to release the sperm. Tyramine and octopamine cause an
increase in the amplitude and
frequency of spermathecal contractions in the locust, L.
migratoria (da Silva and Lange, 2008). In
R. prolixus, RhoprFIRFa and RhoprFMRFa have also been shown to
cause strong contractions in
the bursa, thereby playing a role in oviposition. Proctolin has
also been shown to increase the tone
of bursal contractions in R. prolixus (Chiang et al., 2010).
Therefore, a cocktail of neuroactive
chemicals acting as neurotransmitters, neurohormones and
neuromodulators are important for the
movement of eggs in the reproductive tract.
Significance
As stated earlier, R. prolixus is one of the primary vectors of
Chagas disease. Millions of
people are diagnosed with Chagas disease in Latin America.
Chagas disease does not only cripple
the health and wellbeing of those infected, but also causes
economic losses due to vector control
initiatives. It is known that a mated, regularly fed kissing bug
can lay up to 600 eggs in it’s life
span of 1.5 years (WHO, 2002). This impressive reproductive
capability translates to a massive
number of nymphs arising from a few females. Therefore, it is
very critical to control the
population of the vector to eradicate Chagas disease in epidemic
regions. Octopamine has been
shown to be an important neurochemical associated with the
reproduction and the laying of viable
eggs. Disruption of octopamine signaling leads to reproductive
sterility. The role of octopamine
and tyramine has not been elucidated yet in R. prolixus.
Investigating basic physiology of the
reproductive system will provide information on the role of
octopamine and tyramine signaling
pathways, while studying molecular components will provide
possible targets for octopamine and
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16
tyramine signaling disruption. Perhaps the octopamine and
tyramine GPCRs could be used for
developing novel, species-specific and biostable insecticides
targeting kissing bugs
Thesis objectives
Octopamine and less so tyramine have been implicated in the
regulation of reproductive
processes in many insects such as D. melanogaster, L.
migratoria, Stomoxy calcitrans,
Leucophaea maderae, Gryllus biamaculatus, Nilaparvata lugens,
Periplaneta americana. Most of
the studies available have focused on octopamine’s ability to
modulate contractions of visceral
reproductive muscles. Recently, molecular tools have enabled the
discovery of octopamine
receptors in the female reproductive system of D. melanogaster
and N. lugens. These receptors
were critical for egg-laying. Interestingly, it has been shown
that flies lacking octopamine are
reproductively sterile and retain eggs in the ovary. The purpose
of this thesis is to determine the
role of octopamine and tyramine in modulating the reproductive
system of the adult female R.
prolixus. This will be investigated by first examining the
effects of octopamine and tyramine on
the reproductive musculature and second utilizing the recently
sequenced genome to identify and
functionally characterize octopamine and tyramine GPCRs in the
reproductive system. The aims
of the physiological and molecular approaches are listed
below:
Physiological approach
□ Decipher the effects of octopamine and tyramine on the
rhythmic contractions of the
oviducts and the bursa.
□ Determine the mode of action of octopamine and tyramine at the
oviducts and the bursa.
□ Analyze the effects of injected octopamine and tyramine on
egg-laying.
Molecular approach
□ Isolate and clone octopamine and tyramine GPCRs involved in
signaling reproductive
system.
□ Using online tools, analyze and predict the structural
characteristics of the octopamine and
tyramine receptors.
□ Deorphan the octopamine and the tyramine receptors through a
functional receptor assay.
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17
□ Analyze the spatial expression of octopamine and tyramine
receptor transcripts in the CNS
and in specific reproductive tissues.
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18
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Chapter 2: Octopamine and tyramine regulate the activity of
reproductive
visceral muscles in the adult female blood-feeding bug, Rhodnius
prolixus
Sam Hana, Angela B. Lange
University of Toronto Mississauga, Department of Biology,
Mississauga, ON, Canada L5L1C6.
* Correspondence:
Sam Hana
[email protected]
Keywords: Oviducts, bursa, inhibition, contractions, cyclic
AMP
*** The proceeding chapter is reproduced/adapted with permission
from the Journal of
Experimental Biology.
Octopamine and tyramine regulate the activity of reproductive
visceral muscles in the adult
female blood-feeding bug, Rhodnius prolixus.
Hana, S., and Lange, A. B. (2017).
The Journal of Experimental Biology 220, 1830–1836.
doi:10.1242/jeb.156307.
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24
Abstract
The role of octopamine and tyramine in regulating spontaneous
contractions of
reproductive tissues was examined in the female Rhodnius
prolixus. Octopamine decreased the
amplitude of spontaneous contractions of the oviducts and
reduced RhoprFIRFa-induced
contractions in a dose-dependent manner, whereas tyramine only
reduced the RhoprFIRFa-
induced contractions. Both octopamine and tyramine decreased the
frequency of spontaneous
bursal contractions and completely abolished the contractions at
5×10−7 mol l−1 and above.
Phentolamine, an octopamine receptor antagonist, attenuated the
inhibition induced by octopamine
on the oviducts and the bursa. Octopamine also increased the
levels of cAMP in the oviducts, and
this effect was blocked by phentolamine. Dibutyryl cyclic AMP
mimicked the effects of
octopamine by reducing the frequency of bursal contractions,
suggesting that the octopamine
receptor may act by an Octβ receptor. The tyramine receptor
antagonist yohimbine failed to block
the inhibition of contractions induced by tyramine on the bursa,
suggesting that tyramine may be
acting on the Octβ receptor in the bursa.
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25
Introduction
The biogenic amine octopamine acts as a neurotransmitter,
neuromodulator and
neurohormone in invertebrates (Orchard, 1982). Octopamine and
its precursor tyramine are both
derivatives of the amino acid tyrosine, and octopamine and
tyramine are believed to function
analogously to adrenaline (epinephrine) and noradrenaline
(norepinephrine) in vertebrates
(Roeder, 2005). Thus, tyramine is now considered to be a
neuroactive chemical in its own right,
independent of octopamine (Kononenko et al., 2009; Lange, 2009).
Octopamine and tyramine
regulate diverse physiological and behavioural processes such as
courtship, locomotion, learning
and memory, and reproduction (Avila et al., 2012; Huang et al.,
2016; Roeder, 1999; Selcho et al.,
2012). Female Drosophila melanogaster with mutated tyrosine
decarboxylase show reproductive
sterility due to the lack of octopamine (Cole et al., 2005). In
tyrosine decarboxylase mutant flies,
supplementation with octopamine restored reproductive viability
(Cole et al., 2005). Similarly,
tyramine β-hydroxylase mutant flies that are found to only lack
octopamine are also reproductively
sterile (Monastirioti, 2003; Monastirioti et al., 1996).
Octopamine and tyramine signal via G-
protein coupled receptors (GPCRs), leading to changes in second
messenger levels. The recently
updated receptor classification (Farooqui, 2012) divides the
receptors into Octα-R, Octβ-Rs
(Octβ1-R, Octβ2-R, Octβ3-R), TYR1-R and TYR2-R. In general,
Octβ-Rs lead to elevation of
cAMP while Octα-R and TYR-Rs lead to an increase in Ca2+
(Farooqui, 2012).
The movement of eggs in the reproductive system of Rhodnius
prolixus starts at the ovaries,
the site of egg maturation. Upon ovulation, mature eggs are
released into the oviducts
(Wigglesworth, 1942). Eggs are then guided, via oviductal
peristaltic and phasic contractions, to
the common oviduct, where spermatozoa are released through
spermathecal contractions, leading
to fertilization (Davey, 1958). Fertilized eggs are coated with
secretions from the cement gland
(Lococo and Huebner, 1980). The bursa deposits the fertilized
eggs via strong phasic contractions.
These activities are under the direct control of the central
nervous system (CNS) and branches of
the trunk nerves innervate the reproductive tissues of R.
prolixus (Insausti, 1994). The lateral
oviducts are made up of two layers of visceral muscle, an inner
circular and an outer longitudinal
layer, whilst the bursa is made up of thicker muscle fibres
arranged longitudinally (Sedra and
Lange, 2014). The oviducts and the bursa spontaneously contract
(Sedra and Lange, 2014) but the
site of the intrinsic pacemaker(s) in the reproductive system
has not been identified.
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26
Octopamine and tyramine modulate the myogenic activity of a
variety of visceral muscles
in insects, including tissues of the reproductive system.
Octopamine decreases the basal tonus, and
reduces the amplitude and frequency of neurally evoked
contractions of the lateral oviducts of the
locust Locusta migratoria (Lange and Orchard, 1986). Also,
octopamine has been shown to
decrease the amplitude of proctolin-induced contractions in a
dose-dependent manner (Lange and
Orchard, 1986; Nykamp and Lange, 2000). These effects appear to
be mediated by an Oct/Tyr
receptor shown to be expressed in the oviducts of locusts
(Molaei et al., 2005). In Drosophila and
the stable fly Stomoxys calcitrans, octopamine reduces the
amplitude and frequency of
contractions, and reduces basal tonus of the oviducts in a
dose-dependent manner (Cook and
Wagner, 1992; Middleton et al., 2006; Rodríguez-Valentín et al.,
2006). These physiological
effects could be linked to two receptors: the octopamine
receptor in the mushroom bodies (OAMB)
and Octβ2-R, which have been shown in Drosophila to be expressed
in the epithelial and muscle
cells of the oviducts (Lee et al., 2003; Li et al., 2015; Lim et
al., 2014). These receptors are involved
in ovulation and fertilization of eggs, whereby mutant
constructs of these receptors show
reproductive sterility in females, accumulation of eggs in the
ovary and reduction in the number
of eggs laid (Lee et al., 2003; Li et al., 2015; Lim et al.,
2014). In contrast, octopamine has also
been shown to increase the frequency and the amplitude of
myogenic contractions in the lateral
oviducts of the cricket Gryllus bimaculatus (Tamashiro and
Yoshino, 2014). In the cockroach
Leucophaea maderae, the action of octopamine and tyramine is
unclear; both stimulated oviduct
contractions in some preparations but inhibited oviduct
contractions in other preparations (Cook
et al., 1984). Tyramine decreases the basal tonus and attenuates
proctolin-induced contractions in
locusts (Donini and Lange, 2004); however, tyramine has no
effect on the amplitude of
contractions or basal tonus of the oviducts in D. melanogaster
(Middleton et al., 2006).
The purpose of this study was to determine the role of
octopamine and tyramine in
modulating myogenic contractions of the oviducts and the bursa
of the adult female R. prolixus
and to investigate the mechanism by which octopamine and
tyramine mediate these effects.
Materials and Methods
Animals
Adult R. prolixus Stål 1859 were maintained on a 12 h:12 h
light:dark cycle at
approximately 50% humidity and 28°C. Rhodnius prolixus were fed
defibrinated rabbit's blood
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27
(Hemostat Laboratories, Dixon, CA, USA; supplied by Cedarlane
Laboratories Inc., Burlington,
ON, Canada) once in every instar. Four- to five-week-old unfed
adult females were used for all
experiments.
Chemicals
D,L-Octopamine hydrochloride and tyramine hydrochloride were
made as 10−2 mol l−1
stocks and stored at −20°C. Phentolamine hydrochloride and
dibutyryl cAMP were freshly made
in physiological saline prior to use. Aliquots of AKDNFIRFamide
(RhoprFIRFa, 10−3 mol l−1;
GenScript USA, Inc., Piscataqay, NJ, USA) were stored at −20°C.
Stock solution of yohimbine
was prepared in 95% ethanol; the final percentage of ethanol in
the experimental treatments was
≤0.1%. Physiological saline (NaCl 150 mmol l−1, KCl 8.6 mmol
l−1, CaCl2 2 mmol l−1, NaHCO3 4
mmol l−1, glucose 34 mmol l−1, MgCl2 8.5 mmol l−1, Hepes 5 mmol
l−1, pH 7.2) was prepared in
double distilled water and used to dilute all chemicals. All
chemicals were obtained from Sigma
Aldrich (Oakville, ON, Canada) unless otherwise stated.
Contraction assays
Oviduct bioassay
The wings were cut off and the dorsal cuticle along with the gut
of a female adult R.
prolixus were removed to expose the reproductive system. Using a
fine silk thread, a double knot
was tied at the posterior end of the common oviduct and the
other end of the silk was double
knotted onto the hook of the force displacement signal
transducer (Aksjeselskapet Mikro-
elektronikk, Horten, Norway). The oviducts (lateral and common)
were dissected out and placed
in a Sylgard-coated dish filled with 200 µl of physiological
saline at room temperature. The
anterior end of each lateral oviduct was pinned to the dish with
minutien pins. The signal generated
was amplified, converted into a digital signal by Picoscope 2200
(Pico Technology, St Neots, UK)
and analysed by the PicoLog program (Pico Technology).
To examine the effects of octopamine and tyramine on contraction
of the oviducts, 100 µl
of the bath saline was removed and replaced with 100 µl of
2×10−8 mol l−1 to 2×10−3 mol l−1
octopamine/tyramine. A final volume of 200 µl was maintained at
all times. The tissue was washed
between amine applications. For the inhibitor assays,
phentolamine or yohimbine was mixed with
octopamine or tyramine before application. The amplitude of
three to four contractions (over ∼2
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28
min) was averaged and presented as a percentage relative to
contractions of the tissue in saline
(control). To examine the effects of octopamine or tyramine on a
peptide-induced contraction, 10−6
mol l−1 RhoprFIRFa was used. RhoprFIRFa produced a standard
change in basal tonus which was
then compared with the change in basal tonus produced when the
peptide was applied with the
amine.
Bursa bioassay
The dorsal cuticle was removed followed by the gut, exposing the
reproductive system.
Using a fine silk thread, a double knot was made at the junction
of the bursa and the oviducts. A
cut was made above the double knot and the bursa was left
attached to the ventral cuticle and the
fine silk thread was attached to the force transducer. The bursa
was secured in place by pinning
the ventral cuticle to the Sylgard-coated dish. The amplitude
and the frequency of three to four
contractions were averaged (over ∼2 min) and presented as a
percentage relative to contractions
of the bursa in saline (control). Amines were added to the
preparations as described above for the
oviduct bioassay.
cAMP determination assay
cAMP content in the oviducts of 4- to 6-week-old adult female R.
prolixus was measured.
A total of 50 oviducts were dissected and placed in a dish
containing saline. Two oviducts were
pooled and placed in Eppendorf tubes containing saline. Using a
dispensing pipette, 10 µl of
5×10−3 mol l−1 3-isobutyl-1-methylaxanthine (IBMX) was added to
all tubes followed by the
addition of either octopamine or phentolamine, or both. The
tubes were gently mixed and left to
incubate for 10 min. The reaction was stopped by adding 400 µl
boiling ELISA buffer (Cyclic
AMP ELISA Kit, Cayman Chemical, Ann Arbor, MI, USA). The tubes
were boiled for 10 min
and sonicated for 15 s at output 3 and constant duty cycle with
a Branson Sonifier 250 (VWR,
Mississauga, ON, Canada). The homogenates were centrifuged for
15 min at 13,000 rpm. Two 50
µl samples of supernatant from each tube were assayed for cAMP
with the Cyclic AMP ELISA
Kit (Cayman Chemical) according to the manufacturer's
instructions. The pellets were dissolved
in 100 µl of 1 mol l−1 sodium hydroxide and boiled for 10 min.
The resulting solution was used
for protein determination using Pierce™ BCA Protein Assay kit
(Thermo Fisher Scientific,
Waltham, MA, USA).
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29
Statistical analysis
GraphPad Prism version 5.03 (www.graphpad.com) was used to
create and statistically analyse all
graphs in this paper.
Results
Effect of octopamine and tyramine on lateral oviduct
contractions
The lateral oviducts contracted spontaneously in vitro as a
result of the myogenic activity
of the reproductive musculature (Sedra and Lange, 2014). A
strong phasic contraction was initiated
by one lateral oviduct and was shortly followed by contraction
of the other lateral oviduct. Both
oviducts then relaxed concurrently, causing a single burst (Fig.
1A,B). Twin or triple peaks in the
trace were observed when the two lateral oviducts were not in
sync with each other. Stable
rhythmic activity was maintained for a few hours in
physiological saline. Octopamine reduced the
amplitude of the oviductal contractions in a dose-dependent
manner (Fig. 1C). The amplitude
started to decrease between 10−7 and 10−6 mol l−1 octopamine,
with a significant decrease in
amplitude at 10−5 mol l−1 octopamine (one-way ANOVA followed by
Dunnett's multiple
comparison test compared with the saline group at 100%, P
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30
induced contraction (one-way ANOVA followed by Tukey multiple
comparisons test, P
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31
10−5 mol l−1 also antagonized the effect of 10−5 mol l−1
tyramine in the bursa and restored the
amplitude of contractions to 102.3±6.3% relative to the saline
control.
Yohimbine does not inhibit the effects of tyramine on bursal
contractions
Yohimbine is an α2-adrenergic receptor antagonist known to block
tyramine receptors
(Broeck et al., 1995; Saudou et al., 1990). Yohimbine did not
alter the amplitude of contractions
when applied on its own and did not inhibit the effects of
tyramine on bursal contractions (Fig. 6).
Yohimbine at 10−5 mol l−1 failed to restore bursal contractions
abolished by 10−5 mol l−1
octopamine.
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Figure. 1. The effects of octopamine and tyramine on rhythmic
contractions of the oviducts of an
adult female Rhodnius prolixus. (A) Application of octopamine
(OA, 10−4 mol l−1) inhibits of the
amplitude of contractions. (B) Tyramine (TA, 10−4 mol l−1) does
not affect contractions. The black
bar indicates the period of application of the neurochemical and
the white bar indicates the wash
period. (C) Dose–response curve for the effects of octopamine
and tyramine relative to the
amplitude of contractions in saline prior to the addition of
neurochemicals. Octopamine inhibits
the amplitude of contractions, while tyramine does not affect
contraction amplitude (one-way
ANOVA followed by Dunnett's multiple comparison test; *P
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34
Figure. 2. Octopamine and tyramine effectively antagonize
AKDNFIRFamide (RhoprFIRFa)-
induced contraction of the oviducts of R. prolixus. (A)
Octopamine (10−4 mol l−1) significantly
reduces the amplitude of the RhoprFIRFa (10−6 mol l−1)-induced
contraction. (B) Tyramine (10−4
mol l−1) significantly reduces the RhoprFIRFa (10−6 mol
l−1)-induced contraction. The black bar
indicates the period of application of the neurochemical and the
white bar indicates the wash
period. (C) Inhibition of RhoprFIRFa-induced contraction by
octopamine and tyramine is dose
dependent (one-way ANOVA followed by Tukey multiple comparisons
test; *P
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36
Figure. 3. Octopamine and tyramine abolish the rhythmic
contractions of the bursa in R. prolixus.
(A–D) Application of 10−6 mol l−1 octopamine (A) or tyramine (B)
abolishes contractions of the
bursa. Note that contractions are unchanged at 10−7 mol l−1
octopamine (C) and tyramine (D). The
black bar indicates the period of application of the
neurochemical and the white bar indicates the
wash period. (E) Dose–response curve showing the sudden
abolishment of rhythmic contractions
at concentrations greater than 5×10−7 mol l−1 neurochemical
(n=5–9). (F) Octopamine and
tyramine both significantly decrease the burst frequency
relative to the saline control. (G)
Dibutyryl cAMP reduces the frequency of contractions
significantly at 10−2 mol l−1 relative to the
saline control. (F and G: one-way ANOVA followed by Dunnett's
multiple comparison test;
*P
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38
Figure. 4. Phentolamine blocks the inhibitory effect of
octopamine on rhythmic contractions of
the oviducts. (A) Octopamine (10−4 mol l−1) reduces the
amplitude of spontaneous contraction. (B)
The effect of octopamine (10−4 mol l−1) is inhibited by
application of phentolamine (10−7 mol l−1,
Phen). The black bar indicates the period of application of the
neurochemical and the white bar
indicates the wash period. (C) Phentolamine alone does not
affect the amplitude of contraction.
Phentolamine is capable of reversing the inhibition of oviduct
contraction by octopamine. (D)
Phentolamine attenuates the octopamine-induced rise in cAMP
levels in the oviducts.
Concentrations in C and D are mol l−1. (C and D: one-way ANOVA
followed by Tukey multiple
comparisons test; *P
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39
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40
Figure. 5. Phentolamine blocks abolishment of rhythmic
contractions in the bursa by octopamine.
(A) Octopamine abolishes bursal contractions at 10−5 mol l−1.
(B) Phentolamine (10−5 mol l−1)
blocks the inhibition induced by octopamine (10−5 mol l−1) on
bursal contraction. The black bar
indicates the period of application of the neurochemical and the
white bar indicates the wash
period. (C) Phentolamine at 10−5 mol l−1 does not significantly
increase the amplitude of bursal
contractions when compared with saline. Phentolamine
significantly blocks the inhibitory effect
of octopamine on bursal contractions (concentrations are mol
l−1; one-way ANOVA followed by
Tukey multiple comparisons test; ***P
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41
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42
Figure. 6. Yohimbine fails to block tyramine inhibition of
rhythmic contractions of the bursa. (A)
Tyramine (10−5 mol l−1) inhibits bursa contractions. (B)
Yohimbine (Yhm, 10−5 mol l−1) does not
block the inhibitory effect of tyramine on bursal contractions.
The black bar indicates the period
of application of the neurochemical and the white bar indicates
the wash period. (C) Yohimbine at
10−5 mol l−1 does not block the inhibitory effect of tyramine on
the amplitude of bursal contractions
(one-way ANOVA followed by Tukey multiple comparisons test; not
significant, P>0.05).
Concentrations are mol l−1; means±s.e.m. of n samples noted at
the bottom of each bar; for 10−5
mol l−1 TA, n=4.
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44
Discussion
Octopamine reduced the amplitude of spontaneous rhythmic
contractions in R. prolixus
oviducts. This phenomenon is consistent with results previously
obtained in L. migratoria, D.
melanogaster and S. calcitrans (Cook and Wagner, 1992; Lange and
Orchard, 1986; Middleton et
al., 2006; Rodríguez-Valentín et al., 2006): octopamine reduced
the amplitude of contractions in
S. calcitrans and L. migratoria and reduced neurally evoked
contractions of L. migratoria oviducts
(Cook and Wagner, 1992; Lange and Orchard, 1986). In R.
prolixus, there was no change in
frequency of the oviductal contractions whereas in L. migratoria
and S. calcitrans, octopamine led
to a decrease in basal tonus and frequency in oviduct
contractions. In contrast, in G. bimaculatus,
octopamine increased the amplitude and frequency of rhythmic
contractions in a dose-dependent
manner despite L. migratoria and G. bimaculatus belonging to the
order Orthoptera (Tamashiro
and Yoshino, 2014). In addition, octopamine decreased the
amplitude of the RhoprFIRFa-induced
contraction of R. prolixus oviducts, confirming it as an
inhibitor of oviduct contractions. In locusts,
it was also shown that octopamine reduces proctolin-induced
contractions in the oviducts (Nykamp
and Lange, 2000). Proctolin was also shown to reduce
octopamine-induced cAMP levels in
oviducts, suggesting that the components of octopamine and
proctolin signalling pathways interact
to modulate oviduct contraction (Nykamp and Lange, 2000). The
interaction of RhoprFIRFa and
octopamine is not known, although it is not likely that
octopamine interacts with the RhoprFIRFa
signalling pathway according to a recent study in Drosophila
(Milakovic et al., 2014). Milakovic
et al. (2014) found that FMRFamide-induced muscle contraction is
independent of the well-known
intracellular players such as calmodulin kinase II, IP3, cAMP,
etc.; however, we do not know the
pathway used by RhoprFIRFa in this preparation.
Phentolamine, an α-adrenergic receptor antagonist, is an
effective OctβR antagonist in the
L. migratoria oviduct (Lange and Orchard, 1986; Orchard and
Lange, 1986). Thus, the ability of
phentolamine to block the effects of octopamine on R. prolixus
oviducts suggests that octopamine
is working via an OctβR. This is supported by the fact that
octopamine increases cAMP levels (a
characteristic of OctβRs) in the oviducts, an effect also
blocked by phentolamine (Lange and
Orchard, 1986; Orchard and Lange, 1986). The physiological
implications of these findings in R.
prolixus are that octopamine plays an essential role in the
process of ovulation. Relaxation of the
oviducts would allow the ovary to release more eggs into the
oviducts. In D. melanogaster,
octopamine was found to increase the contractions of the
peritoneal sheath, a contractile meshwork
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45
that surrounds the ovary, and to relax the oviducts, thereby
enabling the release of eggs into the
oviducts (Middleton et al., 2006). Moreover, Octβ2R and OAMB
have been found to be the
receptors associated with the process of ovulation and
fertilization in Drosophila (Lee et al., 2003;
Li et al., 2015; Lim et al., 2014). Deletions in the oamb locus
and mutant constructs of Octβ2R
resulted in accumulation of eggs in the ovary and a significant
decrease in the number of eggs laid
(Lee et al., 2003; Li et al., 2015; Lim et al., 2014).
Similarly, in R. prolixus, octopamine reduced
the amplitude of oviduct contractions, probably by binding to
OctβR, leading to an elevation in
cAMP levels and muscle relaxation.
The process by which tyramine regulates the oviducts seems more
modulatory. Tyramine,
when applied to the oviducts at a wide range of doses did not
elicit any changes in spontaneous
contractions; however, tyramine inhibited RhoprFIRFa-induced
contractions in a dose-dependent
manner. This is not the case in L. migratoria, where tyramine
mimicked octopamine and decreased
the basal tonus and attenuated proctolin-induced contractions
(Donini and Lange, 2004). A
possible explanation for this phenomena in R. prolixus is that
tyramine is co-released with
octopamine; octopamine works on the oviducts to reduce the
amplitude of spontaneous
contractions induced by a pacemaker, whereas octopamine and
tyramine modify the effects of
myogenic stimulators such as RhoprFIRFa.
The effects of octopamine and tyramine on the bursa are similar.
Both biogenic amines
completely abolish contractions of the bursa at high
concentrations. In addition, octopamine and
tyramine at low concentrations decrease the frequency of
contractions. These effects seem to be
mediated by cAMP, as application of the membrane-permeable cAMP
analogue dibutyryl cAMP
decreased the frequency of contractions. Phentolamine
antagonized the effects of octopamine and
tyramine, suggesting that both are likely to work via an OctβR.
Yohimbine did not antagonize the
effects of tyramine and octopamine on the bursa. This suggests
that tyramine acts via the OctβR at
high concentrations, as shown in the locust oviducts and foregut
(Britain, 1990; Donini and Lange,
2004). Further studies are needed to understand why the
contractions are abolished with no
a