A Dissertation entitled
Ectopic Expression in Remodeled C. elegans: A Platform for Target Identification,
Anthelmintic Screening and Receptor Deorphanization
By
Wen Jing Law
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Doctor of Philosophy Degree in Biological Sciences
_________________________________________
Dr. Richard Komuniecki, Committee Chair
_________________________________________
Dr. Bruce Bamber, Committee Member
_________________________________________
Dr. Welivitiya Karunarathne, Committee Member
_________________________________________
Dr. Patricia Komuniecki, Committee Member
_________________________________________
Dr. Scott Molitor, Committee Member
_________________________________________
Dr. Robert Steven, Committee Member
_________________________________________
Dr. Patricia Komuniecki, Dean
College of Graduate Studies
The University of Toledo
May 2016
Copyright 2016, Wen Jing Law
This document is copyrighted material. Under copyright law, no part of this document
may be reproduced without the expressed permission of the author.
An Abstract of
Ectopic expression in remodeled C. elegans: A platform for target identification,
anthelmintic screening and receptor deorphanization
by
Wen Jing Law
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Doctor of Philosophy Degree in Biological Sciences
The University of Toledo
May 2016
Nematode infections cause significant morbidity and have a devastating global economic
impact. Anthelmintic development has been hampered by lack of cost-effective screening
platforms due, in part, to the absence of nematode cell lines, difficulties in maintaining parasitic
nematodes in laboratory, and the costs of in vivo screening. In the present study we exploited the
many advantages of the C. elegans model system and developed a high-throughput screening
platform to identify selective nematode monoamine receptor agonists in genetically-engineered
“chimeric” C. elegans as lead compound for anthelmintic development. Previously, we and
others have demonstrated that exogenous monoamines, such as 5-HT, dopamine and tyramine
(TA), each paralyzed the free-living nematode, C. elegans and, where examined, parasitic
nematodes. Specifically, we have heterologously expressed 5-HT and TA receptors from a variety
of organisms in the motor neurons and body wall muscle of different C. elegans receptor mutants,
at sites yielding robust locomotory phenotypes, including paralysis, upon agonist stimulation.
This approach includes nematode-specific accessory proteins and cuticle, while maintaining the
unique pharmacologies of receptors from individual parasites. Using this approach, we have
identified selective receptor agonists for both the nematode and human 5-HT1 receptors.
Similarly, we have expressed novel nematode monoamine-gated Cl- channels in motor neurons
and body wall muscle and validated their participation in a robust monoamine-mediated paralysis
iii
at both sites. In addition, we have modified the incubation system by varying incubation
conditions and genetic backgrounds to dramatically increase the permeability of the chimeric
animals, allowing much less drug to be used for screening. In fact, using this approach it may be
possible to modify the cuticular permeability of C. elegans to mimic individual parasites species.
Finally, we have used the C. elegans model to tentatively identify the receptor for a
FMRFamide neuropeptide, PF4 (KPNFIRFamide). Previous workers have demonstrated that
PF4 causes a rapid, flaccid paralysis when injected into Ascaris suum and induces a rapid, Cl-dependent hyper-polarization in denervated A. suum muscle strips, suggesting that PF4 opens a
Cl- channel on nematode muscle that has the potential to cause hyper-polarization and flaccid
paralysis and may be a potentially important target for anthelmintic development (Maule et al.,
1995a; Maule et al., 1995b; Holden-Dye et al., 1997; Purcell et al., 2002; Reintiz et al., 2011).
The C. elegans and A. suum flp-1/af26 genes encode peptides similar (or identical) to PF4,
suggesting a potential conservation in both ligand and its receptor. To this end, we identified a
group of sensory-mediated locomotory phenotypes in flp-1 null animals and animals overexpressing flp-1. The C. elegans genome contains a number of genes encoding putative ligandgated Cl- channels. We reasoned that null and XS alleles of the gene encoding the putative PF4
receptor(s) would have phenotypes similar to flp-1 null and XS animals. In addition, flp-1XS
phenotypes should be absent in the receptor null background. This screen identified a putative
receptor meeting these criteria, LGC-50. LGC-50 is highly conserved in both free-living and
parasitic nematodes. As predicted, flp-1 and lgc-50 null phenotypes were identical and overexpressing lgc-50 in the body wall muscle significantly reduced locomotion in wild type, but not
flp-1 null animals. Most importantly, the direct injection of PF4 caused a rapid paralysis in
animals over-expressing LGC-50 in muscle. Together, these genetic data suggest that LGC-50 is
a PF4 receptor; however, PF4 had no effect on Cl- currents of Xenopus oocytes expressing LGC50. A. suum muscle expresses an uncharacterized ligand-gated Cl- channel, with high identity to
iv
the C. elegans LGC-34, based on muscle-specific transcriptome data, so we also examined lgc-34
null animals for flp-1 phenotypes. Surprisingly, lgc-34 null and over-expression phenotypes were
identical to those observed in flp-1 and lgc-50 null and over-expressing animals. Most
importantly, lgc-50 over-expression phenotypes were absent in lgc-34 null animals, suggesting
that LGC-50 and LGC-34 might both be subunits of a PF4-gated channel. These studies are
continuing to characterize the ligand-specificity of the putative LGC-50/LGC-34 heteromeric
channel.
These studies highlight the power of the C. elegans model system for target identification
and anthelmintic screening. Certainly, any observations from “chimeric” C. elegans will
ultimately need to be verified in the target parasite, but these results validate the utility and
versatility of this “dual systems” approach, where observations from free-living nematodes are
translated to the parasites. Although the present study is focused on monoamine receptors, the
screening protocol and approaches that we have developed have the potential to be useful in the
characterization a wide range of additional anthelmintic targets.
v
Acknowledgements
To my advisor, Dr. Richard Komuniecki, for his constant guidance and insights
into the works presented in this thesis, and most importantly the super-human patience he
has shown toward me and my idiosyncrasies all these years.
Heartfelt thanks to Dr. Vera Hapiak and Dr. Gareth Harris for showing me the
“worm trade”. Also, many thanks to Amanda Ortega for all the microinjection and
transgenic lines used in this project, and to Mitch and Toby, for putting up with this
oddball.
Lastly, I would like to thank Drs. Bruce Bamber, Welivitiya Karunarathne,
Patricia Komuniecki, Scott Molitor and Robert Steven for taking the troubles to be my
committee member when Christmas is just round the corner.
vi
Contents
Abstract …………………………………………………………………………….… iii
Acknowledgements ……………………………………………………………..….. vi
Contents ……………………………………………………………………………... vii
List of Figures ……………………………………………………………………..… x
List of Abbreviations …………………………………………………………...... xiii
List of Symbols ……………………………………………………………………. xiv
Preface ………………………………………………………………………………… 1
1 Significance
1.1
Part 1 ……………………………………………………………….......... 3
1.2
Part 2 …………………………………………………………………….. 8
2 Materials & Methods
2.1 Strains and Reagents …………………………………………………………… 13
2.2 Fusion PCR and Transgenic lines ……………………………………………… 14
2.3 Paralysis assay …………………………………………………………………. 15
vii
2.4 Octanol avoidance assay ……………………………………………………….. 16
2.5 Peptide injection assay …………………………………………………………. 16
2.6 Locomotory assay …………………………………………………………….... 16
2.7 Accession numbers …………………………………………………………….. 17
3 Results
3.1 Part 1
Rationale ………………………………………………………………….…… 18
3.1.1
5-HT inhibits locomotion in 5-HT receptor null animals expressing 5-HT1like receptors in the AIB interneurons or cholinergic motor neurons….. 20
3.1.2
Use of heterologous expression for agonist identification ……………... 25
3.1.3
Identification of agonists with potential selectivity for a nematode 5-HT1like receptor ………………………………………………………...….. 29
3.1.4
The activation of monoamine-gated Cl- channels in cholinergic motor
neurons or body wall muscles causes locomotory paralysis …………… 30
3.1.5
The inhibition of AIB signaling causes “locomotory confusion” and
paralysis …...…………………………………………………………… 33
3.1.6
The activation of an excitatory GPCR in cholinergic motor neurons also
causes locomotory paralysis …...………………………………………. 35
3.2 Part 2
Rationale ……………………………………………………………………….. 37
3.2.1
flp-1 and lgc-50 null animals are hyper-responsive to 30% 1-octanol in the
absence of food ………………………………………………………… 38
3.2.2
flp-1 and lgc-50 over-expressors are stimulated by salt and not inhibited
by TA and OA ………………………………………………………….. 40
3.2.3
LGC-50 functions in ASE, ASH, ASI and/or AWC sensory neurons, based
on cell-specific RNAi and the rescue of lgc-50 null animals ………….. 43
viii
3.2.4
The direct injection of PF4 into animals over-expressing lgc-50 in body
wall muscle causes paralysis …………………………………………… 48
3.2.5
Animals over-expressing lgc-50 in body wall muscle have reduced
mobility in the presence of food ……………………………………….. 49
3.2.6
LGC-50 may form a heterologous channel with other ligand-gated ion
channel subunits ……………………………………………………….. 51
4 Discussion ………………………………………………………………………… 56
References ………………………………………………………………………….. 68
ix
List of Figures
1.1.1
Phylogenetic relationship among monoamine G-protein coupled receptors
(GPCRs) from C. elegans and their orthologues in both free-living and parasitic
nematodes. ………………………………………………………………………. 6
1.1.2
Putative orthologues of various cys-loop receptors are highly conserved in both
parasitic and free-living nematodes. …………………………………………….. 7
3.1.1
C. elegans mutants with increased cuticular permeability are hyper-sensitive to 5HT-dependent paralysis. ……………………………………………………….. 23
3.1.2
The 5-HT/SER-4-dependent inhibition of either the AIB interneurons or
cholinergic motor neurons causes locomotory paralysis. ……………………… 25
3.1.3
5-HT and 5-HT receptor agonists selectively paralyze C. elegans 5-HT receptor
mutant animals expressing nematode, insect or human 5-HT1-like receptors in the
cholinergic motor neurons. …………………………………………………….. 27
3.1.4 PAPP-dependent paralysis requires the 5-HT1-like receptor, SER-4 and the D1like dopamine receptor, DOP-3. ……………………………………………….. 28
3.1.5 Identification of compounds with selectivity for nematode 5-HT1A receptors as
potential lead compounds for potential anthelmintic development. …………… 30
x
3.1.6
Exogenous monoamines paralyze C. elegans expressing monoamine-gated Clchannels in either cholinergic motor neurons or body wall muscles. ………….. 32
3.1.7
Inhibiting signaling from the two AIB interneurons causes “locomotory
confusion” and paralysis. ………………………………………………………. 34
3.1.8
5-HT paralyzes 5-HT receptor quintuple null animals expressing either the Gαqcoupled 5-HT receptor, SER-7b or the Gαs-coupled 5-HT receptor, SER-1a in the
cholinergic motor neurons. …………………………………………………….. 36
3.2.1 Aversive responses to 30% 1-octanol are more rapid in flp-1 and lgc-50 null
mutants. ………………………………………………………………………… 39
3.2.2
Aversive responses to 30% -octanol are more rapid after the RNAi knockdown of
lgc-50 or flp-1 in wild-type animals. …………………………………………… 40
3.2.3
Aversive responses in animals over-expressing lgc-50 or flp-1 are stimulated by
small increases in the Cl- concentration. ……………………………………….. 41
3.2.4
Aversive responses in animals over-expressing lgc-50 or flp-1 are stimulated by
NaCl and are not inhibited by tyramine or octopamine. ……………………….. 43
3.2.5
An Plgc-50::lgc-50(+)::GFP transgene is expressed in head muscle, as well as a
number of head and tail neurons, including the ASI sensory neurons and the
ventral cord motor neurons. ……………………………………………………. 45
3.2.6
Aversive responses to 30% 1-octanol are more rapid after the neuron-selective
RNAi knockdown of lgc-50 in the ASI, ASH, ASE AWC and/or AWB sensory
xi
neurons in wild type animals, mimicking the more rapid aversive phenotype
observed in lgc-50 null animals. ……………………………………………….. 46
3.2.7
The neuron-selective expression of lgc-50 in ASI, ASH, ASE AWC and/or AWB
sensory neurons in lgc-50 null animals can rescue the hyper-responsiveness (5 s)
to 30% 1-octanol observed in lgc-50 null animals to wild type level (10 s). ….. 47
3.2.8
The direct injection of PF4 (100 µM) into the pseudocoelom of animals overexpressing lgc-50 in the body wall muscle (Pmyo-3) causes rapid onset of
paralysis. ……………………………………………………………………….. 49
3.2.9
Animals over-expressing lgc-50 in body wall muscle (Pmyo-3::lgc-50::GFP
transgene) move more slowly than wild-type animals in the presence of food
(OP50). ………………………………………………………………………… 50
3.2.10 Many cys-loop, ligand-gated Cl- channel null mutants respond more rapidly to
30% 1-octanol than wild-type animals. ………………………………………... 52
3.2.11 LGC-34 and LGC-47 may form a heteromeric peptide-gated ion channel with
LGC-50. ………………………………………………………………………... 53
3.2.12 Aversive responses in animals over-expressing lgc-50 or lgc-34 are stimulated by
NaCl and not inhibited by tyramine or octopamine. …………………………… 54
xii
List of Abbreviations
ser-4 ………….. G-protein (Gαo) coupled serotonin receptor
mod-1 ………… serotonin-gated chloride channel
lgc-55 ………… tyramine-gated chloride channel
ser-1…………... G-protein (Gαq) coupled serotonin receptor
ser-7…………... G-protein (Gαs) coupled serotonin receptor
5-HT quint ……. ser-5;ser-4;mod-1;ser-7;ser-1 quintuple null
SER-4 quad …... ser-5;mod-1;ser-7;ser-1 quadruple null
TA quad ………. tyra-2;tyra-3;ser-2;lgc-55 quadruple null
Hco …………… Haemonchus contortus
5-HT ………….. serotonin
TA …………….. tyramine
DA ……………. dopamine
GFP …………… green fluorescence protein
RFP …………… red fluorescence protein
8-OH-DPAT …... 8-hydroxy-2-(di-n-propylamino)tetralin
PAPP ………….. p-amino-phenethyl-m-trifluoromethylphenyl piperazine
His …………….. histamine
ACh …………… acetylcholine
DCBP …………. 1,2-dibromo-3-chloropropane
GPCR …………. G-protein coupled receptor
xiii
List of Symbols
α ……………….. Alpha
β ……………….. Beta
xiv
Preface
Parasitic nematodes are a major threat to human health and socioeconomic wellbeing around the world, especially in developing countries (Awasthi & Bundy, 2007;
Hotez et al., 2008; Wani et al., 2010). Unlike tuberculosis and malaria, parasitic
nematode infections usually result in morbidity, rather than the mortality that tends to be
the focus of media and policy makers. Morbidity and the resulting loss of disabilityadjusted life years (DALYs), particularly in children and the younger population, is
especially devastating in poorer regions, as they hinder any development efforts to lift
these regions out of poverty (Hotez et al., 2008; Hotez & Kamath, 2009; Brooker, 2010).
Despite the dire situation, the development of new human anthelmintics is a low priority
for pharmaceutical companies because of a lack of economic incentives, given that the
target market cannot afford the new drug at prices high enough to recoup research
investment and generate profit (Kaplan, 2004; Zhang et al., 2010).
Another less reported facet of parasitic nematode disease is its impact on
agriculture, especially with respect to large-scale industrialized livestock and crop
production. For example, parasitic nematodes infect livestock and major crops (corn and
soybeans) and cause billions in economic losses yearly in the US alone (Jones et al.
2013). Indeed, the marketing and development of almost all current anthelmintics is
sustained and driven by demands from agricultural sectors, as most human anthelmintic
1
development and distribution are dependent on donations and funding from governments,
pharmaceutical companies and non-governmental organizations (Waller, 1999; Kaplan,
2004; Geary et al., 2010; Zhang et al., 2010). However, the banning of nematicides in
crop production due to environmental toxicity, and the continued reliance on the few
“traditional” anthelmintics (levamisole, ivermectin etc.) and their derivatives has given
rise to increasing levels of resistant in parasitic nematodes of both humans and livestock
(Reynoldson et al., 1997; Bain, 1999; Albonico et al., 2002; Kaplan, 2004;
Wolstenholme et al., 2004). Furthermore, some parasitic nematode diseases, like
filariasis and onchocerciasis, for which no effective chemotherapy has been developed,
remain as major causes of DALYS in affected areas. Current anti-filarial drugs target the
mosquito-infective microfilaria, not the adult parasite, and require mass drug
administration over an extended period to interrupt transmission (Keating et al., 2014).
New drugs, new drug targets and new, more effective screening protocols are desperately
needed in all settings.
2
Chapter 1
Significance
1.1 Part 1
Most anthelmintics in use today act as agonists at key receptors and cause
paralysis by interfering with muscle contraction and/or locomotion (Martin, 1985; Geary
et al., 1993; Sheriff et al., 2002; Martin et al., 2012). Since receptor “activation” is
essential for anthelmintic activity, receptor knockout is not necessarily the “gold
standard” for target validation; in fact knockout may not be lethal. Five molecular targets
have been used for drug discovery, two nicotinic cholinergic receptor subunits
(tetrahydropyrimidines/ imidathiazoles and amino-acetonitriles), glutamate-/GABA-gated
Cl- channels (macrocyclic lactones and piperazine, respectively) and Ca++-gated K+
channels (emodepside) (Martin, 1985; Geary et al., 1993; Sheriff et al., 2002; Martin et
al., 2012). Importantly, anthelmintics targeting each of these molecular targets are active
in the free-living nematode, Caenorhabditis elegans and our understanding of their
modes of action has, in large part, resulted from our ability to genetically manipulate their
putative targets in receptive C. elegans mutant backgrounds (Holden-Dye et al., 2012;
Krucken et al., 2012; Miltsch et al., 2013; Hernando & Bouzat, 2014). Importantly, the
identification of new targets has been limited by the lack of useful information about the
3
identity, function and localization of the additional receptors regulating muscle contraction and locomotion. In addition to identifying new targets, we also need new highthroughput screening protocols that preserve the unique pharmacologies of the receptors
from the different parasites and maintain a nematode-specific context that includes the
cuticle and appropriate accessory proteins, especially given that no nematode cells lines
are available and that the parasites themselves are extremely difficult and expensive to
culture.
In the present study, we have developed a novel, heterologous, ectopic overexpression approach to provide a unique nematode screening platform for selective
agonist identification, exploiting the unique experimental advantages of the C. elegans
model system. Previously, we and others have demonstrated that exogenous
monoamines, such as serotonin (5-HT), dopamine (DA) and tyramine (TA), each
paralyze C. elegans and, where examined, parasitic nematodes (Ranganathan et al., 2000;
Hobson et al., 2006; Hapiak et al., 2009; Gurel et al., 2012; Chase et al., 2004;
McDonald et al., 2007; Allen et al., 2011; Rex et al., 2004; Donnelly et al., 2013; Reinitz
& Stretton, 1996; Masler, 2007; Beech et al., 2013). In each case, the key C. elegans
receptors mediating this locomotory inhibition have been identified and functionally
localized, with each operating at a different level within the locomotory circuit: 5-HT
requires the Gαo-coupled 5-HT1-like GPCR, SER-4 and the 5-HT gated Cl- channel ,
MOD-1 in a few key interneurons, including the two AIBs, DA the Gαo-coupled DA
GPCR DOP-3 in ventral cord GABAergic and cholinergic motor neurons and TA the
TA-gated Cl- channel, LGC-55, in head muscle and the Gαo/Gq-coupled TA GPCRs
4
SER-2, TYRA-2 and TYRA-3 in as yet unidentified interneurons (Hapiak et al., 2009;
Allen et al., 2011; Donnelly et al., 2013).
Importantly, these C. elegans receptors have clear orthologues in many medically/
agriculturally important parasitic nematodes, as shown in Figure 1.1.1 & 1.1.2. This
conservation potentially allows the characterization of these receptors in the relatively
well established C. elegans model system. Furthermore, C. elegans appears to be a highly
promiscuous expression platform, able to functionally express receptors from diverse
origins including Drosophila and humans, while also maintaining the ligand selectivity/
specificity of these receptors, potentially allowing most if not all parasitic nematode
receptors to be characterized in this convenient model organism (Law et al., 2015;
Komuniecki, Mills & Oakes, unpublished).
5
Figure 1.1.1. Phylogenetic relationship among monoamine G-protein coupled
receptors (GPCRs) from C. elegans and their orthologues in both free-living and
parasitic nematodes.
Predicted protein sequences were aligned using ClustalW and an unrooted tree was
constructed using a Neighbourhood-Joining method in MEGA5. A. suum sequences were
based on the A. suum draft genome (Jex et al., 2011). Sequences for C. elegans (Ce), C.
brenneri, C. briggsae, C. remanei, B. malayi, L. loa and H. contortus were obtained from
GenBank. Accession numbers for C. elegans are: Ce dop-1: NP_001024577.1, Ce dop-2:
NP_001024048.1, Ce dop-3: NP_001024908.2, Ce dop-4: NP_508238.2, Ce dop-5:
NP_505884.1, Ce dop-6: NP_508739.3, CE octr-1: NP_001024569.1, Ce ser-1:
NP_001024728.1, Ce ser-2: NP_001024339.1, Ce ser-3: NP_491954.1, Ce ser-4:
NP_497452.1, Ce ser-5: NP_492273.2, Ce ser-6: NP_741350.1, Ce ser-7: NP_741730.1,
Ce tyra-2: NP_001033537.1, Ce tyra-3: NP_001024805.1. Species and receptor
orthologues are color-coded: red for parasitic species and blue/ black for free-living
species.
6
Figure 1.1.2. Putative orthologues of various cys-loop receptors are highly conserved
in both parasitic and free-living nematodes.
This phlylogenetic tree was generated using the “blastp” & “Cobalt” software available at
http://blast.ncbi.nlm.nih.gov/Blast.cgi. A. suum (As) cys-loop receptors sequences were
based on the A. suum draft genome (2011) by Jex et al. Sequences for C. elegans (Ce),
C. brenneri (Cbr), C. briggsae (Cb), C. remanei (Cr), B. malayi (Bm), L. loa (Ll) and H.
contortus (Hc) were obtained from GenBank. The accession numbers for lgc-50: Ce NP_498637.3, Cbr - EGT56575.1, Cb - XP_002642590.1, Cr - XP_003095901.1, Bm XP_001894837.1, Ll - XP_003141833.1; acc-1: Ce - NP_501715.1, Cbr - EGT33480.1,
Cb - XP_002647842.1, Cr - XP_003094757.1, As - ADY44658.1, Bm XP_001899286.1; acc-3: Ce - NP_508810.2, Cbr - EGT30171.1, Cb - XP_002644586.1,
Cr - XP_003117949.1, Ll - XP_003142307.1; acc-4: Ce - NP_499789.1, Cbr EGT60165.1, Cb - XP_002643140.1, Cr - XP_003102137.1, Ll - XP_003142764.1; lgc46: Ce - NP_497338.2, Cbr - EGT57109.1, Cb - XP_002640981.1, Cr XP_003102299.1, Bm - XP_001900347.1; lgc-55: Ce - NP_507870.2, Cbr EGT49133.1, Cb - XP_002638506.1, Cr - XP_003094349.1, Hc - ACZ57924.1, Ll XP_003141089.1; mod-1: Ce - NP_741580.1, Cbr - EGT39436.1, Cb XP_002637422.1, Cr - XP_003097605.1, As - ADY43724.1. Species name and their
respective cys-loop channels are color-coded: parasitic nematodes in red and free-living
nematodes are in blue/ black.
We have previously constructed quintuple 5-HT receptor null C. elegans (5-HT
quint) that do not express any previously identified 5-HT receptors and do not respond to
7
exogenous 5-HT in a range of behavioral assays, to identify essential roles for the Gαocoupled 5-HT1-like SER-4 and the unique 5-HT-gated Cl- channel, MOD-1 in 5-HTdependent locomotory paralysis (Hobson et al., 2006; Hapiak et al., 2009). Importantly,
SER-4 agonists appear to function as anthelmintics in vivo and have been used to clear
Haemonchus contortus infections from gerbils (Smith et al., 2003; White et al., 2007). In
the present study, we ectopically expressed SER-4 and MOD-1 orthologues from
parasitic nematodes, insects and humans in either the cholinergic motor neurons or body
wall muscles of quintuple C. elegans 5-HT receptor null animals, on the assumption that
agonist-dependent receptor activation at these sites will cause robust locomotory
phenotypes and paralysis that can be readily adapted to high-throughput screening
protocols. For example, the activation of a ligand-gated Cl- channel in body wall muscles
would be predicted to hyper-polarize the muscle and significantly inhibit locomotion,
while the activation of a Cl- channel or Gαo-coupled GPCR on the cholinergic motor
neurons would be predicted to significantly inhibit ACh release from the motor neurons
and inhibit both muscle contraction and thus, locomotion.
1.2 Part 2
Ion channels in the neuromuscular system of parasitic nematodes have always
been the preferred target of human/animal anthelmintic development, because the
therapeutic consequences are usually both rapid and robust. In fact, most popular
anthelmintics, such as ivermectin (glutamate-gated Cl- channel), levamisole (nAch-gated
Na+ channel) and piperazine (GABA-gated Cl- channel) activate neuromuscular ionchannels, causing spastic (ivermectin & levamisole) or flaccid (piperazine) paralysis in
8
parasite muscles, ultimately inhibiting a wide range of key behaviors, including
locomotion, egg-laying and pharyngeal pumping, depending on the location of their
respective receptors (Martin, 1985; Geary et al., 1993; Gill et al., 1998; Sheriff et al.,
2002). Importantly, resistance has arisen rapidly to all classes of anthelmintics, including
most recently ivermectin (Lustigman & Carter, 2007). Therefore, the search for and
development of new, more effective anthelmintics that activate these, and as yet
unidentified ion channels is worthwhile goal.
Nematodes express a wide range of neuropeptides and neuropeptide receptors.
For example, C. elegans has over 100 predicted neuropeptide genes that encode over 250
distinct peptides (Li et al. 1999a; Li & Kim, 2010). Of these, 40 encode insulin-like
peptides, 30 FMRFamide-like peptides and the remainder NLP peptides (noninsulin/non-FMRFamide). In addition, the C. elegans genome encodes over 60 predicted
neuropeptide receptors. Importantly, many of these neuropeptide-encoding genes and
neuropeptide receptors appear to be essential for locomotory decision-making, based on a
range of locomotory phenotypes in individual neuropeptide null animals and most have
clear orthologues in the genomes of parasitic nematodes (Maule et al., 1995a; Geary et
al., 1999; Li et al. 2005; Reinitz et al., 2011). Over the years, many research groups have
identified a number of myoactive neuropeptides that showed promise as new
peptidomimetic anthelmintics, but the general cost-effectiveness of mainstream
anthelmintics has prevented pharmaceutical companies from further developing these
drug candidates (Kaplan, 2004; Geary, 2005). This endeavor is further hampered by the
lack of bioinformatic tools available for parasitic nematodes. Indeed, sequencing projects
for genomes of major parasites like Brugia malayi, Ascaris suum and Haemonchus
9
contortus have been only completed recently (Ghedin et al., 2007; Jex et al., 2011; Laing
et al., 2013; Schwarz et al., 2013). However, with resistance to mainstream
anthelmintics targeting classical neurotransmitter-gated ion channels on the rise, focus on
re-exploring the possibility of neuropeptides and their receptors as novel anthelmintic
targets has been renewed, as highlighted in several recent publications as well as the
discovery and adoption of emodepside, which targtes Ca++-gated K+ channels, as a novel
anthelmintic (Martin & Robertson, 2010; Reinitz et al., 2011; McVeigh et al., 2012). To
date, with the exception of a few FMRFamide-gated Na+ channels identified in mollusks
and cnidarians, all of the previously characterized neuropeptide receptors are G-protein
coupled (Kubiak et al., 2003a; Thompson et al., 2003; Lingueglia et al., 2006; Furukawa
et al., 2006; Durrnagel et al., 2010).
The FMRFamide neuropeptide, PF4 (KPNFIRFamide), was first identified in the
free-living nematode, Panagrellus redivivius and causes a rapid, flaccid paralysis when
injected into the pseudocoelom of adult A. suum. PF4 also induces a rapid, Cl--dependent
hyper-polarization and flaccid paralysis of isolated A. suum muscle strips, suggesting that
the PF4 receptor may be a potentially potent new target for anthelmintic development
(Maule et al., 1995a; Maule et al., 1995b; Holden-Dye et al., 1997; Purcell et al., 2002;
Reintiz et al., 2011). Similar peptides have also been identified in a wide-variety of freeliving and parasitic nematodes (Cowden & Stretton, 1995; Maule et al., 1996; Kim & Li,
2004; McVeigh et al., 2005; McVeigh et al., 2006). For example, in A. suum, the afp-11
gene encodes multiple (12) peptides, including PF4 and a similar peptide
KPNFLRFamide (AF26), many of which, including AF26, also cause flaccid paralysis
when injected into pseudocoelom of A. suum (Reinitz et al., 2000; Reinitz et al., 2011).
10
In C. elegans, a flp-1-encoded peptide, KPNFLRFamide, is nearly identical to PF4/AF26,
both structurally and in amino acid composition, and also causes a rapid relaxation of
isolated A. suum muscle strips (Rosoff et al., 1992; Rosoff et al., 1993; Maule et al.,
1995a). Perhaps more importantly, PF4/AF26 also cause a rapid, Cl--dependent, hyperpolarization of denervated A. suum muscle strips, suggesting that PF4 directly gates Clchannels located post-synaptically on the muscle, leading to hyper-polarization and
flaccid paralysis (Maule et al., 1995a). These observations make the PF4/AF26 receptor
a potentially useful target for anthelmintic development.
The C. elegans genome encodes a number of predicted cys-loop, ligand-gated Clchannel subunits that are activated by glutamate, acetylcholine and GABA, each of which
has homologues in mammals (Jones & Sattelle, 2008). However, in contrast to mammals
and higher eukaryotes, channels activated physiologically by the monoamines 5-HT, TA,
and DA, and choline have also been identified, making the ligand diversity of the
nematode Cl- channels much greater that their mammalian counterparts (Ranganathan et
al., 2000; Pirri et al., 2009; Ringstad et al., 2009). All members of the cys-loop, ligandgated ion channel family share the same basic structure, with five subunits surrounding a
selective, ion-conducting pore. Each subunit is made up of 4 membrane-spanning αhelices (M1-4), an extracellular domain containing the characteristic cys-loop and a
relatively large intracellular domain between M3 and M4. The 2nd α-helix (M2) is the
channel-lining domain and contains the ion-selective motifs, such as the PARS motif of
the Cl- channels. The N-terminal extracellular domain is responsible for ligand-binding,
whereas the intercellular loop between M3 and M4 is involved in receptor trafficking,
assembly and modulation, as well as portals to allow ions entering or leaving the cell.
11
M4 also interacts with lipids and neurosteroids to modulate channel activity (Carswell et
al., 2015). Upon ligand binding to the extracellular loop, the channel undergoes
conformational changes to allow the correct ion species to flow down electro-chemical
gradients into or out of the cell (Thompson et al., 2010). The different possible subunit
combinations (either subunit species or ratio in channel composition) of heteromeric cysloop ion channels can contribute to pharmacological differences with regard to ligand
specificity and sensitivity, adding to the complexity in an already diverse group of
receptors and contributing to the difficulties in identifying the exact composition of a
channel for a particular ligand, as demonstrated by nACh receptor (Boulin et al., 2008;
Williamson et al., 2009; Durrnagel et al., 2010; Boulin et al., 2011; Bennett et al., 2012).
Furthermore, the possibility exists that each monomeric/ heteromeric channel could be
activated by more than one endogenous ligand. However, these complexities offer a
notable advantage, because while many of these ligand-gated channels are conserved
among nematode species, even with mammals, the ligand selectivity and sensitivity
generated by subtle differences in receptor structure, together with the ligand diversity
within the nematode phylum provide a fertile ground for the development of highly
specific anthelmintics (Williamson et al., 2009).
12
Chapter 2
Materials & Methods
2.1 Strains and Reagents
bus-8 (e2968), bus-16 (e2802), bus-17 (e2800), flp-1 (ok2811), lgc-50 (tm3712),
lgc-34 (gk532), lgc-40 (n4545), acc-1 (tm3268), acc-2 (tm3219 & ok2216), acc-3
(ok3450), acc-4 (ok2371), lgc-46 (ok2949), lgc-47 (ok2963 & ok3016), lgc-49 (tm5183),
lgc-51 (tm4318), lgc-52 (tm4268), lgc-53 (n4330), lgc-54 (T15B7.16) and lgc-55 (n4331)
were obtained from Caenorhabditis Genetics Center (CGC). ser-5 (tm2654);ser-4
(ok512);mod-1 (ok103);ser-7 (tm1325) ser-1 (ok345) (5-HT quint), ser-5 (tm2654);mod1 (ok103);ser-7 (tm1325) ser-1 (ok345) (SER-4 quad) and lgc-55 (tm2913);tyra-3
(ok325) tyra-2 (tm1846) ser-2 (pk1357) (TA quad) were generated as described
previously (Hobson et al., 2006; Hapiak et al., 2009). All strains were maintained on
NGM plates with OP50 at 16°C.
The cDNA clone of Drosophila melanogaster 5-HT1A (RE57708) was ordered
from the Drosophila Genomics Resource Center (DGRC), the cDNA clone Human
HTR1A (MGC: 167873; clone ID: 9020250) from GE Healthcare Dharmacon Inc. and
cDNA clones of Haemonchus contortus (Hco) lgc-55 and mod-1 orthologues were kindly
provided by Dr. Sean Forrester (Rao et al., 2010; Beech et al., 2013). The unc-17ß
13
promoter, RM#621p, was obtained from Dr. James Rand. The integrated AIB::HisCl1 in
N2 (cx15457) animals were a kind gift from Dr. Cornelia Bargmann (Pokala et al., 2014).
Serotonin (5-HT) (H7752-25G), tyramine (TA) (T2879-25G), 8-OH DPAT
(H141-25MG), sumatriptan succinate (S1198-10MG), PAPP (S009-25MG), histamine
(H7250-5G) and octopamine (OA) (O0250-5G) were purchased from Sigma Life
Sciences. Stock solutions (50 mM) of 5-HT, TA, 8-OH-DPAT, sumatriptan and
histamine were made up in distilled water, PAPP in 100% ethanol. Compounds for
screening (CD3-238, -257, -276, -531, -664, -717, -718, -946, -947, -980, -984, BK-4-15,
CT-3-38, MOFLIPP & MOMIPP were kind gift from Dr. Paul Erhardt and The
University of Toledo Center for Drug Design and Development (CD3). Stock solutions
(200 mM) were made up in 100% DMSO. The constituent of for nematode growth
media (NGM), potassium phosphate monobasic (KH2PO4; P285-3), sodium chloride
(NaCl; S271-3), calcium chloride dehydrate (CaCl2.2H2O; C79-500), magnesium sulfate
heptahydrate (MgSO4.7H2O; BP213-1), tryptone (BP1421-2) and agar (DF0812071)
were purchased from Thermo Fisher Scientific Inc., cholesterol (C3045-5G) purchased
from Sigma Life Science.
2.2 Fusion PCR and Transgenic lines
All transgenic constructs were created by overlap fusion PCR (Hobert, 2002). All
transgenes contain a GFP marker (with unc-54 3′-UTR) at the 3‟-end. PCR products
from multiple reactions were pooled and co-injected with coelomocyte-RFP screening
marker into the appropriate null backgrounds (Mello & Fire, 1995). Once generated,
14
transgenic animals were frozen in liquid nitrogen and thawed fresh weekly for assay.
Multiple transgenic lines from each construct were examined.
2.3 Paralysis assay
Fresh agar plates (without NaCl, KH2PO4, MgSO4, CaCl2, tryptone and
cholesterol) containing 5-HT, TA, PAPP, sumatriptan or 8-OH DPAT at desired
concentrations were made daily. For assays involving bus mutants, fresh NGM agar
plates (with NaCl, KH2PO4, MgSO4, CaCl2, tryptone and cholesterol) containing 5-HT
were used for all assays. For assays with AIB::HisCl1 (cx15457) animals, freshly poured
NGM agar or agar only plates containing 10 mM and 2 mM histamine were used. NGM
agar plates were prepared as described in WormBook (Stiernagle, 2006).
For all paralysis assays, well-fed, transgenic young adults expressing RFP
screening markers were picked 2 hrs prior to assay and maintained on NGM plates with
E. coli OP50. For assay, 10 animals are transferred to assay plates (agar only for all
assays and NGM agar for assays with bus mutants) containing the appropriate drug and
motility was assessed at intervals of 5 min for 30 min. Experiments with sumatriptan
were carried out for 60 min, with motility assessed every 5 min. All assays were
conducted in the absence of food, i.e. OP50. Animals that moved less than 1 body bend/
20 s were counted as paralyzed. Each transgenic line was assayed at least 3 times with 10
animals/assay for each agonist concentration. Data is presented as % paralyzed ± SE
over drug exposure time (min). Dose-response curves and EC50s were then generated
using a variable slope nonlinear regression model with GraphPad Prism 6 software. Drug
concentrations were log10-transformed prior to analysis.
15
2.4 Octanol avoidance assay
Well-fed hermaphrodite forth-stage (L4) C. elegans larvae were picked 24 h prior
to assay and incubated overnight at 20°C. Fresh nematode growth media (NGM) plates
were prepared on the day of assay. Additional 4 mM of either 5-HT, TA, OA or NaCl
were added as required. Aversive responses were examined as described in Chao et al.,
2004 and are presented as the time taken to initiate backward locomotion after the
presentation of 30% 1-octanol on a hair in front of a forward moving animal. Each strain
was assayed for at least 3 times with 25 animals/assay. Data is presented as mean ± SE
and analyzed by two-tailed Student‟s t-test.
2.5 Peptide injection assay
KPNFLRFa (100 µM in M9) were injected into the pseudocoelom of wild type
(N2), lgc-50 and wild type animals over-expressing lgc-50 in the body wall muscle
(Pmyo-3). Animals were examined on fresh NGM plates with no addition at 5 min
intervals for 20 min. Animals that moved less than 1 body bend/ 20 s were counted as
paralyzed. Data are presented as mean  SE and analyzed by a two-tailed Student‟s t test.
2.6 Locomotory assay
Well-fed hermaphrodite forth-stage (L4) C. elegans larvae were picked 24 h prior
to assay and incubated overnight at 20°C. Fresh nematode growth media (NGM) plates
were prepared on the day of assay. OP50 were added after agar solidified and were
allowed to dry out prior to assay. Animals were examined for locomotion (no. of body
bends/ 20 s) and frequency of pauses in 3 min. A pause is tentatively defined as
16
temporary cessation of forward locomotion before resumption of locomotion along the
original trajectory. Data are presented as mean  SE and analyzed by a two-tailed
Student‟s t test.
2.7 Accession numbers
The accession numbers of the proteins involved in our study are C. elegans SER-4
(accession no. NP_497452), C. elegans LGC-55 (accession no. NP_507870), C. elegans
MOD-1 (accession no. CCD72364), D. melanogaster 5-HT1A (accession no.
NM_166322.2), D. melanogaster HisCl1 (accession no. Q9VGI0), human HTR1A
(accession no. BC136263), H. contortus LGC-55 (accession no. ACZ57924.1) and H.
contortus MOD-1 (accession no. ADM53350.1).
17
Chapter 3
Results
3.1 Part 1
Rationale: The monoamines, 5-HT, DA and TA, each dramatically inhibit locomotion in
C. elegans when applied exogenously at concentrations high enough to overcome the
permeability barrier of the nematode cuticle, ultimately resulting in paralysis (Hapiak et
al., 2009; Gurel et al., 2012; Donnelly et al., 2013). Using the C. elegans model, the
receptors involved in monoamine-dependent locomotory inhibition have been identified
and localized (Ranganathan et al., 2000; Hobson et al., 2006; Hapiak et al., 2009; Gurel
et al., 2012; Chase et al., 2004; McDonald et al., 2007; Allen et al., 2011; Rex et al.,
2004; Donnelly et al., 2013). Interestingly, the key receptors involved in 5-HT, DA and
TA inhibition each function at a different level in the locomotory circuit with 5-HTdependent paralysis requiring the expression of the Gαo-coupled, 5-HT1-like receptor,
SER-4, and the 5-HT-gated Cl- channel, MOD-1 in a limited number of interneurons,
including the two AIBs (Hapiak et al., 2009; Gurel et al., 2012). Unfortunately, since
nematode cell lines are not available and the maintenance of parasitic nematodes outside
their hosts is problematic, screening platforms for anti-nematodal activity have been
limited and do not usually incorporate the nematode cuticle or potentially important
nematode accessory proteins.
18
The present study was designed to develop a screening platform for nematode
monoamine receptor agonists in “chimeric” genetically-engineered C. elegans by
heterologously expressing 5-HT and TA receptors at sites likely to yield robust
locomotory phenotypes upon agonist stimulation. Previously, many investigators have
rescued a range of behaviors in C. elegans null animals with the expression of proteins
from the parasites, validating this approach (Kaminsky et al., 2008; Crisford et al., 2011;
Welz et al., 2011). We chose to examine locomotion as an endpoint for heterologous,
ectopic expression, as the neurons and circuits modulating locomotion in C. elegans and
parasitic nematodes appear to be highly conserved, can be readily adapted to established
screening assays, and have always been the primary target for the majority of existing
anthelmintics. Specifically, we expressed 1) Gαo-coupled, 5-HT1-like receptors, or 5-HT/
TA-gated Cl- channels in the cholinergic motor neurons of C. elegans mutants lacking
any known 5-HT or TA receptors, respectively on the assumption that robust agonistdependent Gαo signaling or potential hyper-polarization, respectively, would dramatically
inhibit ACh release and locomotion. 2) 5-HT or TA-gated Cl- channels in body muscle
of C. elegans mutants lacking any known 5-HT or TA receptors, respectively, on the
assumption that agonist-dependent muscle hyper-polarization would cause paralysis. 3)
Gαs/Gαq-coupled 5-HT receptors in cholinergic motor neurons of C. elegans mutants
lacking any known 5-HT receptors, on the assumption that robust agonist-dependent
Gαs/Gαq signaling would stimulate over-release of ACh, resulting in over-active muscle
contraction and hence spastic paralysis. As noted below, all three hypotheses have been
confirmed.
19
3.1.1. 5-HT inhibits locomotion in 5-HT receptor null animals
expressing 5-HT1-like receptors in the AIB interneurons or cholinergic
motor neurons
The role of the C. elegans 5-HT1-like receptor, SER-4, in 5-HT-dependent
paralysis is well documented (Hobson et al., 2006; Hapiak et al., 2009; Gurel et al.,
2012; Komuniecki et al., 2012). Indeed, the utility of the H. contortus SER-4 orthologue,
5-HT1Hco as an anthelmintic target has been validated previously both in vivo and in vitro
(Smith et al., 2003; White et al., 2007). Locomotion in C. elegans has been assessed
previously using a number of different assays, many of which can be readily adapted for
screening (Ramot et al., 2008; Smout et al., 2010; Wang & Wang, 2013; Buckingham &
Sattelle, 2009; Chen et al., 2011; Carr et al., 2011). For example, automated thrashing
assays allow thousands of compounds to be easily screened per day (Buckingham &
Sattelle, 2009). Monoamine-dependent locomotory inhibition and paralysis has been
quantified on agar plates (sinusoidal body bends) and in liquid medium (C-shaped
“swimming”), containing either M9 buffer or water (Ranganathan et al., 2000; Rex et al.,
2004; McDonald et al., 2007; Hapiak et al., 2009; Gurel et al., 2012; Donnelly et al.,
2013). The permeability of the C. elegans cuticle appears to vary depending on
incubation conditions, with much less 5-HT apparently required in water, than in saltcontaining media (M9), possibly because of an increased cuticular permeability under
hypotonic conditions (Gurel et al., 2012).
Previously, we assayed locomotion under standard C. elegans culture conditions
on NGM agar plates. Under these conditions, 15 mM 5-HT initiated a rapid paralysis in
wild type animals, and ser-5;mod-1;ser-7 ser-1 quadruple null (SER-4 quad) animals
20
(Hapiak et al., 2009; Komuniecki et al., 2012). As predicted, 5-HT had no effect on
locomotion in 5-HT quint animals that lack all previously identified 5-HT receptors
(Figure 3.1.1A-B) (Hapiak et al., 2009). This 5-HT-dependent paralysis was not the
classical spastic paralysis associated with cholinergic agonists, such as levamisole, or the
flaccid paralysis associated with glutamatergic agonists, such as ivermectin, but instead
appeared to result more from “locomotory confusion,” with animals unable to effectively
integrate conflicting sensory inputs to initiate and sustain forward/ backward locomotion.
The C. elegans cuticle appears to be more impermeable than those of some of the
parasitic nematodes (Ho et al., 1992; Page & Johnstone, 2007; Ruiz-Lancheros et al.,
2011). Therefore, since the concentration of 5-HT required for maximal paralysis was
quite high (15 mM) in these short term assays, presumably to overcome cuticular
permeability, we re-assayed these animals under hypotonic conditions on agar plates
without salt (non-NGM) (Figure 3.1.1C-D). Attempts to repeat published data from
others on 5-HT paralysis in water were unsuccessful, as majority of the animals burst
soon (within 5 min) after exposure to water (Gurel et al., 2012). However, in a hypotonic
environment (agar alone without NGM), much lower concentrations of 5-HT were
required for inhibition of wild type animals, with 1 mM 5-HT yielding 50% paralysis
after 10 min exposure (EC50 about 0.4 mM) (Figure 3.1.1C-D).
In addition to hypotonic incubation, we also examined 5-HT-dependent paralysis
in a number of C. elegans mutants that exhibit increased cuticular permeability. For
example, the Hodgkin group previously identified a series of bus mutants that exhibit
increased cuticular permeability that have been hypothesized to be excellent vehicles for
small molecule screening (Partridge et al., 2008). Indeed, as noted in Figure 3.1.1E-F,
21
many of the bus mutants are hyper-sensitive to 5-HT-dependent paralysis, even under
isotonic assay conditions (on NGM agar plates). For example, bus-17 mutants are
acutely paralyzed after 10 min on 5-HT with an EC50 of about 0.24 mM, which is
substantially lower than that observed in wild-type animals incubated under the same
conditions (EC50 = 11.5 mM) (Figure 3.1.1F). These results suggest that these mutants
might be useful for agonist identification, especially when only limited amounts of
compound are available. Indeed, it may even be possible to select mutants that exhibit
cuticular permeabilities that mimic those of individual parasites. Unfortunately, these
mutants are also sensitive to hypotonicity and burst under the hypotonic conditions used
in the present study, so that they could not be used in combination with hypotonicity to
further increase sensitivity (data not shown). Therefore, unless specified, hypotonic
conditions were used to assay the transgenic animals described below.
22
Figure 3.1.1. C. elegans mutants with increased cuticular permeability are hypersensitive to 5-HT-dependent paralysis.
A-B. Paralysis of wild type and mutant C. elegans on NGM agar plates. A. Wild type
animals examined for 5-HT-dependent paralysis as outlined in Methods. Data are
presented as mean ± SE (n=3). 5-HT quint animals were not paralyzed by 5-HT at the
23
concentrations examined (data not shown). B. Dose-response curves for 5-HT-dependent
paralysis on NGM plates at 10 min exposure for wild type and 5-HT quint animals. C-D.
Paralysis of wild type and mutant C. elegans on non-NGM agar (hypotonic) plates. C.
Wild type animals were examined for 5-HT-dependent paralysis as outlined in Methods.
Data are presented as mean ± SE (n=3). 5-HT quint animals were not paralyzed by 5-HT
at the concentrations examined (data not shown). D. Dose-response curves for 5-HTdependent paralysis in hypotonic conditions at 15 min exposure for wild type and 5-HT
quint animals. E-F. 5-HT-dependent paralysis of wild type and mutant C. elegans on
NGM agar plates. E. 5-HT (0.25 mM)-dependent paralysis of wild-type, bus-8 (e2968),
bus-16 (e2802) and bus-17 (e2800) mutants. Data are presented as mean ± SE (n=3). F.
Dose-response curves for 5-HT-dependent paralysis at 10 min exposure for wild type and
bus mutants.
A ser-4::gfp transgene is expressed in a limited number of neurons, including the
AIBs (Gurel et al., 2012). Therefore, SER-4::GFP was specifically expressed in either
the AIB interneurons (Pnpr-9) or ectopically, in the cholinergic motor neurons (Punc17β) of the 5-HT quint. Expression was confirmed by GFP fluorescence (Figure 3.1.2A).
As predicted, 5-HT quint animals expressing SER-4 in either the AIBs or cholinergic
motor neurons were rapidly paralyzed by 5-HT (Figure 3.1.2B). Interestingly, on 5-HT,
although 5-HT quint animals expressing SER-4 in the AIBs alone moved only
infrequently, they initiated backward locomotion for a short distance when prodded with
a blunt platinum wire at the tail, suggesting that they were probably unable to process
conflicting locomotory signals, as hypothesized above. In contrast, animals expressing
SER-4 in the cholinergic motor neurons were fully paralyzed and did not move when
prodded.
24
Figure 3.1.2. The 5-HT/SER-4-dependent inhibition of either the AIB interneurons
or cholinergic motor neurons causes locomotory paralysis.
A. Confocal images of 5-HT quint expressing SER-4::GFP in the AIB interneurons
(Pnpr-9)(A1) or cholinergic motor neurons (Punc-17β)(A2). GFP fluorescence (A2) or
GFP fluorescence overlaid on DIC image (A1). B. Paralysis of wild type, mutant and
transgenic C. elegans on hypotonic, non-NGM agar plates. Wild type, quadruple 5-HT
receptor null animals expressing only SER-4 (SER-4 quad) or 5-HT quint expressing the
C. elegans 5-HT1-like receptor, SER-4, in either the cholinergic motor neurons (Punc17β) or the two AIB interneurons (Pnpr-9) were examined for 5-HT (1 mM)-dependent
paralysis as outlined in Methods. Data are presented as mean ± SE (n=3).
3.1.2. Use of heterologous expression for agonist identification
To demonstrate the utility of this screening approach, the Drosophila 5-HT1
orthologue (5HT1A) or the human 5-HT-1A receptor (HTR1A) were also expressed
specifically in the cholinergic motor neurons (Punc-17β) of 5-HT quint animals.
Locomotion in animals from both transgenic lines was dramatically inhibited by
exogenous 5-HT, demonstrating that the receptors were functionally expressed (Figure
3.1.3A). To demonstrate the specificity of these chimeric C. elegans for agonist
identification, we examined the effect of 8-hydroxy-2-(di-n-propylamino)tetralin (8-OHDPAT), a subtype-selective agonist for the human 5-HT1A receptor, sumatriptan
succinate, a selective mammalian 5-HT1B/D agonist, and p-amino-phenethyl-m-
25
trifluoromethylphenyl piperazine (PAPP). As predicted, 8-OH-DPAT rapidly paralyzed
the 5-HT quint animals expressing the human 5-HT1A receptor (Figure 3.1.3B). In
contrast, 8-OH-DPAT, even at 2 mM, had no effect on locomotion of 5-HT quint animals
expressing either Drosophila or C. elegans 5-HT1 receptor orthologues, suggesting the
conservation of ligand-receptor specificity in chimeric C. elegans (Figure 3.1.3B).
Sumatriptan, at low concentrations, is a selective mammalian 5-HT1B/D agonist, and,
indeed in the present study, sumatriptan was much less effective than 8-OH-DPAT in
initiating paralysis (Razzaque et al., 1999). For example, 0.5 mM sumatriptan had no
effect on locomotion in either wild type or transgenic animals expressing 5-HT1A receptor
orthologues in cholinergic motor neurons (data not shown) and, even at higher
concentrations, failed to fully paralyze animals expressing the human 5-HT1A receptor.
In addition, although animals expressing the human 5-HT1A receptor responded to
increased sumatriptan concentrations more rapidly, these locomotory effects were
transient and reduced dramatically after 25 min, presumably due to receptor
desensitization (Figure 3.1.3C). In contrast, paralysis increased with prolonged
sumatriptan exposure in animals expressing either the C. elegans or Drosophila
receptors, demonstrating kinetic differences between the orthologous receptors.
26
Figure 3.1.3. 5-HT and 5-HT receptor agonists selectively paralyze C. elegans 5-HT
receptor mutant animals expressing nematode, insect or human 5-HT1-like
receptors in the cholinergic motor neurons.
A-C. Paralysis of wild type, mutant and transgenic C. elegans on hypotonic, non-NGM
agar plates. A. 5-HT (1 mM)-dependent paralysis of 5-HT quint animals expressing
either C. elegans 5-HT1-like (SER-4), Drosophila 5-HT1-like, or human 5-HT1A receptor
in cholinergic motor neurons (Punc-17β). Data are presented as mean ± SE (n = 3). B. 8OH-DPAT (2 mM)-dependent paralysis of 5-HT quint animals expressing either C.
elegans 5-HT1-like (SER-4), Drosophila 5-HT1-like, or human 5-HT1A receptor in
cholinergic motor neurons (Punc-17β). Data are presented as mean ± SE (n=3). C.
Sumatriptan (1 mM)-dependent paralysis of wild type, 5-HT quint animals expressing
either C. elegans 5-HT1-like (SER-4), Drosophila 5-HT1-like, or human 5-HT1A receptor
in cholinergic motor neurons (Punc-17β). Data are presented as mean ± SE (n=3).
PAPP, a high affinity agonist for the H. contortus 5-HT1-like receptor, paralyzes
H. contortus L3s in vitro and clears experimental H. contortus infections from gerbils
(Smith et al., 2003; White et al., 2007). As predicted, PAPP initiated a rapid paralysis in
wild type animals (EC50 = 0.37 mM) and, even more rapidly, in 5-HT quint animals
expressing the C. elegans SER-4 in the cholinergic motor neurons (EC50 = 0.17 mM),
supporting the previous identification of PAPP as a 5-HT1-like receptor agonist (Figure
3.1.4A-B). In contrast, and somewhat surprisingly, at higher concentrations (≥0.5 mM),
PAPP also paralyzed 5-HT quint animals (EC50 = 0.68 mM) that were unaffected by 5-
27
HT, suggesting that, in addition to acting as a 5-HT1-like receptor (SER-4) agonist, PAPP
also acted at second target(s) (Figure 3.1.4A-B). Since exogenous TA and DA also
paralyze C. elegans, we surmised that, at higher concentrations, PAPP might be
activating additional monoamine receptors. DA-dependent paralysis requires the
expression of the Gαo-coupled DA receptor, DOP-3 in the cholinergic motor neurons
(Chase et al., 2004). Therefore, dop-3 expression was knocked down in the 5-HT quint
animals using dop-3 RNAi driven by the dop-3 promoter. As noted in Figure 3.1.4C,
dop-3 RNAi knockdown in this background significantly reduced PAPP-dependent
paralysis, suggesting that DOP-3 is a secondary PAPP target. Screening is in progress to
identify additional target(s). Together, these data highlight the utility of this approach in
preliminary drug screening and suggest that it may also be useful for the identification of
nematode-specific agonists.
Figure 3.1.4. PAPP-dependent paralysis requires the 5-HT1-like receptor, SER-4 and
the D1-like dopamine receptor, DOP-3.
A-C. Paralysis of wild type, mutant and transgenic C. elegans on hypotonic non-NGM
agar plates. A. PAPP (0.5 mM)-dependent paralysis of wild-type, 5-HT quint and 5-HT
quint animals expressing SER-4 in the cholinergic motor neurons (Punc-17β). Data are
presented as mean ± SE (n=3). B. Dose-response curves for PAPP-dependent paralysis at
28
15 min exposure for wild type, 5-HT quint and 5-HT quint animals expressing SER-4 in
the cholinergic motor neurons (Punc-17β). C. PAPP (0.5 mM)-dependent paralysis of 5HT quint and 5-HT quint animals expressing Pdop-3::dop-3 RNAi. Data are presented as
mean ± SE (n=3). „*‟ p≤0.001, significantly different from 5-HT quint animals assayed
under identical conditions.
3.1.3. Identification of agonists with potential selectivity for a nematode
5-HT1-like receptor
To demonstrate the utility of our screening approach, we proceeded with a
preliminary screen of 17 compounds with structural similarity with 5-HT, provided by
The University of Toledo Center for Drug Discovery and Design (CD3). Among the 17
compounds examined, CD3-718, CD3-664, CD3-980, CD3-984, CD3-276, CD3-947 and
CD3-946 exhibited comparatively higher selectivity for the nematode 5-HT1-like receptor
(SER-4), paralyzing 5-HT quint animals expressing SER-4 in the cholinergic motor
neurons (Punc-17β) more robustly than 5-HT quint animals expressing the human 5-HT1like receptor (HTR-1A) (Figure 3.1.5). Indeed, CD3-718 and CD3-664 appeared to be
especially specific to the nematode receptor, paralyzing only the 5-HT quint animals
expressing SER-4 at concentrations as low as 0.5 mM. Although far from being
conclusive, this result further demonstrates the utility of the current approach as a simple,
rapid way to identify potential compounds for further investigation.
29
Figure 3.1.5. Identification of compounds with selectivity for nematode 5-HT1A
receptors as potential lead compounds for potential anthelmintic development.
Paralysis of mutant and transgenic C. elegans on non-NGM agar plates. Wild type, 5-HT
quint and 5-HT quint animals expressing either human 5-HT1A receptor (HTR1A) or C.
elegans 5-HT1-like receptor, SER-4 in the cholinergic motor neurons (Punc-17β) were
examined for 5-HT-like compound (0.5 mM)-dependent paralysis on non-NGM agar
plates after 15 min drug exposure. Data are presented as mean ± SE (n=3).
3.1.4. The activation of monoamine-gated Cl- channels in cholinergic
motor neurons or body wall muscles causes locomotory paralysis
Nematodes also express a unique family of monoamine-gated Cl- channels that
appear to be highly conserved within the phylum, including the C. elegans 5-HT- and
TA-gated Cl- channels, MOD-1 and LGC-55, that play key roles in 5-HT- and TAdependent muscle paralysis, respectively. The C. elegans MOD-1 and its H. contortus
orthologue were expressed directly in either cholinergic motor neurons (Punc-17β) or
body wall muscles (Pmyo-3) of 5-HT quint animals and 5-HT-dependent paralysis was
assayed as described above. Muscle expression was confirmed by GFP fluorescence
30
(Figure 3.1.6A). As previously noted, 5-HT had no effect on locomotion in 5-HT quint
animals, but rapidly paralyzed the 5-HT quint animals expressing either the C. elegans
MOD-1 in the cholinergic motor neurons or the H. contortus MOD-1 orthologue in
cholinergic motor neurons or body wall muscle, with EC50s of about 0.3 mM, 0.2 mM
and 0.2 mM, respectively (Figure 3.1.6B-C). Interestingly, 5-HT-dependent paralysis
was more rapid in the transgenic animals expressing MOD-1 orthologues in the
cholinergic motor neurons than in wild type animals.
Similarly, LGC-55 was expressed in the body wall muscles (Pmyo-3) or its H.
contortus orthologue in the cholinergic motor neurons (Punc-17β) of lgc-55;tyra-3 tyra-2
ser-2 quadruple TA receptor null (TA quad) animals. TA quad animals lack all
previously identified TA receptors and fail to respond to TA in a range of behavioral
assays, including locomotion. As predicted, TA had no effect on locomotion in the TA
quad animals, but significantly inhibited locomotion in TA quad animals expressing
either C. elegans LGC-55 in body wall muscles or H. contortus LGC-55 orthologue in
cholinergic motor neurons, each with EC50 of about 0.1 mM (Figure 3.1.6D-E).
Together, these data suggest that monoaminergic activation of these Cl- channels hyperpolarizes either the cholinergic motor neurons or body wall muscles and inhibits muscle
contraction, as well as highlighting the utility of chimeric C. elegans as a functional
expression platform to identify ligand-gated Cl- channels agonists for use as
anthelmintics.
31
Figure 3.1.6. Exogenous monoamines paralyze C. elegans expressing monoaminegated Cl- channels in either cholinergic motor neurons or body wall muscles.
A. Confocal image of 5-HT quint animals expressing H. contortus MOD-1::GFP in body
wall muscles (Pmyo-3). GFP-fluorescence image. B-D. Paralysis of wild type, mutant
and transgenic C. elegans on non-NGM agar plates. B. 5-HT (0.5 mM)-dependent
paralysis of wild type, 5-HT quint and 5-HT quint animals expressing either the C.
elegans or H. contortus MOD-1 orthologues in the cholinergic motor neurons (Punc-17β)
or the H. contortus MOD-1 orthologue in body wall muscle (Pmyo-3). Data are presented
as mean ± SE (n=4). C. Tyramine (1 mM)-dependent paralysis of wild type, TA quad
and TA quad animals expressing either the C. elegans LGG-55 in body wall muscle
(Pmyo-3) or the H. contortus LGC-55 orthologue in the cholinergic motor neurons (Punc17β). Data are presented as mean ± SE (n=3). D. Dose-response curves for TAdependent paralysis at 15 min exposure for wild type, TA quad and TA quad animals
32
expressing either LGC-55 in the body wall muscles (Pmyo-3), or H. contortus LGC-55
orthologue in cholinergic motor neurons (Punc-17β).
3.1.5. The inhibition of AIB signaling causes “locomotory confusion”
and paralysis
Our results suggest that inhibiting AIB signaling by the expression of a Gαocoupled 5-HT receptor in the AIBs of the 5-HT quint can cause “locomotory confusion”
and subsequent paralysis (Figure 3.1.2B). Similarly, the AIB-specific expression (Pinx1) of the 5-HT-gated Cl- channel, MOD-1 can also cause paralysis (Figure 3.1.7A). In
contrast, ablation of the AIBs does not cause paralysis (Gray et al., 2005; Piggott et al.,
2011). Interestingly, the activation of a Drosophila histamine-gated Cl- channel (HisCl1)
expressed ectopically in the AIBs (cx15457) with 2 mM exogenous histamine caused
AIB hyper-polarization and locomotory phenotypes, but not paralysis (Pokala et al.,
2014). In contrast, increasing the histamine concentration to 10 mM caused paralysis that
persisted for up to 24 hrs in the presence of histamine (Pokala et al., 2014). Similarly, in
the present study, 2 mM histamine did not cause paralysis in wild type animals or in
transgenic animals expressing HisCl1 in the AIBs (cx15457) on NGM plates (Figure
3.1.7B). However, 2 mM histamine caused significance paralysis under the modified
hypotonic assay conditions used in the present study or when the histamine concentration
was raised to 10 mM on NGM plates (Figure 3.1.7B-C). Since the ablation of the AIBs
does not cause paralysis, these results support our previous hypothesis that the partial
inhibition of AIB signaling by partial hyper-polarization or the activation of Gαo
signaling causes an imbalance in the locomotory circuit that results in a state of decisionmaking “confusion,” an inability to execute and sustain unidirectional movement and
33
ultimately, in cessation of locomotion (paralysis). Theoretically, any ligand that
selectively unbalances AIB signaling has the potential to yield a similar locomotory
phenotype and its target a potential site for anthelmintic development.
Figure 3.1.7. Inhibiting signaling from the two AIB interneurons causes
“locomotory confusion” and paralysis.
A-C. Paralysis of wild type, mutant and transgenic C. elegans on either NGM or nonNGM agar plates. A. 5-HT quint and 5-HT quint animals expressing MOD-1 in the AIBs
(Pinx-1) were examined for 5-HT (1 mM)-dependent paralysis on non-NGM agar plates,
as outlined in Methods. Data are presented as mean ± SE (n=3). B and C. Wild type
animals expressing HisCl1 in the AIBs (cx15457) were examined for histamine (2 or 10
mM)-dependent paralysis on NGM (B) and non-NGM (C) agar plates. Wild type
animals were not paralyzed by histamine (data not shown). Data are presented as mean ±
SE (n=3).
34
3.1.6. The activation of an excitatory GPCR in cholinergic motor
neurons also causes locomotory paralysis
In contrast to the inhibitory effects of ectopically-expressed Gαo-coupled, 5-HT1like receptors and 5-HT/TA-gated Cl- channels on locomotion described above, 5-HT
also initiated paralysis in 5-HT quint animals over-expressing excitatory SER-1a (Gαqcoupled) and SER-7b (Gαs-coupled) in the cholinergic motor neurons (Punc-17β),
presumably due to the over-release of ACh causing over-contraction in body wall
muscles and hence spastic paralysis (Figure 3.1.8A-B). Indeed, 5-HT quint animals
expressing the SER-1a in the cholinergic motor neurons appeared to be significantly
more sensitive to exogenous 5-HT than the other 5-HT GPCRs, with >50% of animals
paralyzed within 10 min of exposure to 0.5 mM 5-HT (as opposed to the 1 mM 5-HT
usually required for optimal paralysis for the other cholinergic motor neuron-expressed 5HT GPCRs). That paralysis can be achieved via two totally opposing cellular signaling
pathways further highlighted the versatility of this ectopic expression approach for the
identification of potential species-specific monoamine receptor agonists.
35
Figure 3.1.8. 5-HT paralyzes 5-HT receptor quintuple null animals expressing either
Gαs-coupled 5-HT receptor, SER-7b or Gαq-coupled 5-HT receptor, SER-1a in the
cholinergic motor neurons.
A-B. Paralysis of wild type, mutant and transgenic C. elegans on non-NGM agar plates.
A. Wild type, 5-HT quint and 5-HT quint animals expressing SER-7b in the cholinergic
motor neurons (Punc-17β) were examined for 5-HT (1 mM)-dependent paralysis on nonNGM agar plates. B. Wild type, 5-HT quint and 5-HT quint animals expressing SER-1a
in the cholinergic motor neurons (Punc-17β) were examined for 5-HT (0.5 mM)dependent paralysis on non-NGM agar plates. Data are presented as mean ± SE (n=3).
36
3.2 Part 2
Rationale: The FMRFamide neuropeptide PF4 (KPNFIRFamide) causes a rapid, flaccid
paralysis when injected into the pseudocoelom of adult A. suum and induces a rapid, Cl-dependent hyper-polarization and flaccid paralysis of A. suum muscle, suggesting that the
PF4 receptor may be a potentially important new target for anthelmintic development
(Maule et al., 1995b; Holden-Dye et al., 1997; Purcell et al., 2002; Reintiz et al., 2011).
The C. elegans flp-1 and A. suum af26 genes encode peptides identical/similar to PF4, as
well as several other neuropeptides, so we have exploited the molecular genetics of the C.
elegans model system to tentatively identify the PF4 receptor(s) and indeed, receptors for
the other FLP-1 peptides. To this end, we identified a group of sensory-mediated
locomotory phenotypes in flp-1 null animals and animals over-expressing flp-1. The C.
elegans genome contains a number of genes encoding putative ligand-gated cys-loop Clchannels. We reasoned that nulls and over-expressors of the gene encoding the putative
PF4 and/or FLP-1 receptor(s) would have phenotypes similar to flp-1 nulls and overexpressors, respectively. This screen identified genes encoding three putative cys-loop
receptors meeting these criteria, LGC-34, LGC-47 and LGC-50. lgc-34, lgc-47 and lgc50 encode a cys-loop Cl- channel subunits that are highly conserved in both free-living
and parasitic nematodes. The present study was designed to characterize these potential
PF4 receptors to provide potentially new anthelmintic targets.
37
3.2.1. flp-1 and lgc-50 null animals are hyper-responsive to 30% 1octanol in the absence of food
Both flp-1 and lgc-50 null animals are hyper-responsive to 30% 1-octanol in the
absence of food, initiating an aversive response to the repellant in about 5s, in contrast to
wild type (N2) animals that respond in about 10 s when assayed under identical
conditions (Figure 3.2.1). As predicted, this hyper-responsive phenotype in lgc-50 null
animals can be rescued to the by expression of a full length lgc-50 genomic construct
with a 5 kb promoter upstream of the ATG (Figure 3.2.1). Similarly, a full length flp-1
genomic construct with a 3 kb promoter upstream of the ATG site is capable of rescuing
the flp-1 null phenotype (Figure 3.2.1). As predicted, RNAi knockdown of lgc-50 or flp1 in N2 animals reproduced the hyper-responsive phenotype observed in the
corresponding null animals (Figure 3.2.2). Many previously reported flp-1 null
phenotypes appear to result from the deletion of daf-10 that is encoded within the flp-1
gene and not the deletion of flp-1 itself, so this RNAi knockdown data is important for
confirming the flp-1 phenotype and may also explain why other reported flp-1 phenotypes
were not observed in lgc-50 null animals (Ailion & Thomas, 2001).
38
Figure 3.2.1. Aversive responses to 30% 1-octanol are more rapid in flp-1 and lgc-50
null mutants.
Animals were examined on fresh NGM plates without addition for their ability to respond
to 30% 1-octanol, as we have described previously. Data are presented as mean  SE and
analyzed by a two-tailed Student‟s t test. „*‟p<0.001, significantly different from wildtype animals assayed under identical conditions.
39
Figure 3.2.2. Aversive responses to 30% -octanol are more rapid after the RNAi
knockdown of lgc-50 or flp-1 in wild-type animals.
Animals were examined on fresh NGM plates without addition for their ability to respond
to 30% 1-octanol, as we have described previously. Data are presented as mean  SE and
analyzed by a two-tailed Student‟s t test. „*‟p<0.001, significantly different from wildtype animals assayed under identical conditions.
3.2.2. flp-1 and lgc-50 over-expressors are stimulated by salt and not
inhibited by TA and OA
Over-expressing either flp-1 or lgc-50 in wild type animals had no effect of
aversive responses to 30% 1-octanol either on or off food (data not shown). In contrast,
animals over-expressing either flp-1 or lgc-50 were hyper-responsive to 30% 1-octanol
when exposed to a 4 mM increase in NaCl concentration on NGM plates (total 52 mM
NaCl) (Figure 3.2.3). Indeed, increasing the NaCl concentration by as little as 0.5 mM
induced this hyper-response to 30% 1-octanol in both lgc-50 and flp-1 over-expressors
(data not shown). In addition, salt dependent phenotype appeared to involve Cl-, not Na+.
40
For example, lgc-50/ flp-1 over-expressors on plates containing 4 mM ammonium
chloride (NH4Cl), mimicked animals on plates containg additonal 4 mM NaCl and
exhibited hyper-responses to 30% 1-octanol. In contrast, lgc-50/ flp-1 over-expressors on
plates containing 4 mM sodium acetate (C2H3NaO2) exhbited wild-type responses to 30%
1-octanol (Figure 3.2.3).
Figure 3.2.3. Aversive responses in animals over-expressing lgc-50 or flp-1 are
stimulated by small increases in the Cl- concentration.
Animals were examined on fresh NGM plates with additional 4 mM NaCl, 4 mM NH4Cl
or 4 mM C2H3NaO2 for their ability to respond to 30% 1-octanol, as we have described
previously. Data are presented as mean  SE and analyzed by a two-tailed Student‟s t
test. „*‟p<0.001, significantly different from wild-type animals assayed under identical
conditions.
Serotonin induces a hyper-response to 30% 1-octanol in wild-type animals,
identical to that observed above for lgc-50/ flp-1 over-expressors on salt (Chao et al.,
2004; Harris et al., 2009; Harris et al., 2010; Harris et al., 2011; Mills et al., 2012;
41
Hapiak et al., 2013). TA or OA inhibited the hyper-response to 30% 1-octanol observed
in 1) wild-type animals in the presence of 5-HT and 2) flp-1 or lgc-50 null animals
(Harris et al., 2010; Mills et al., 2012; Hapiak et al., 2013) (Figure 3.2.4). In contrast,
TA and OA had no effect on the hyper-response to 30% 1-octanol observed in animals
over-expressing either flp-1 or lgc-50 on 4 mM NaCl (Figure 3.2.4). Therefore, to
demonstrate that the hyper-response to 30% 1-octanol observed in animals overexpressing flp-1 on 4 mM NaCl was dependent on lgc-50, we over-expressed flp-1 in an
lgc-50 null background.
As predicted, since lgc-50 null animals alone exhibit a hyper-
response to 30% 1-octanol, animals over-expressing flp-1 in an lgc-50 null background
also exhibited a hyper-response to 30% 1-octanol (Figure 3.2.4). However, in contrast to
animals expressing flp-1 in a wild-type background that were not inhibited by TA or OA,
animals expressing flp-1 in an lgc-50 null background were inhibited by TA or OA,
suggesting that the more rapid locomotion in the animals expressing flp-1 in an lgc-50
null background was due to the absence of lgc-50 and NOT flp-1-dependent lgc-50
signaling.
42
Figure 3.2.4. Aversive responses in animals over-expressing lgc-50 or flp-1 are
stimulated by NaCl and are not inhibited by tyramine or octopamine.
Animals were examined on fresh NGM plates with additional 4 mM NaCl, 4 mM TA or
4 mM OA for their ability to respond to 30% 1-octanol, as we have described previously.
Data are presented as mean  SE and analyzed by a two-tailed Student‟s t test.
„*‟p<0.001, significantly different from wild-type animals assayed under identical
conditions.
3.2.3. LGC-50 functions in ASE, ASH, ASI and/or AWC sensory
neurons, based on cell-specific RNAi and the rescue of lgc-50 null
animals
An lgc-50 translational fusion with GFP (Plgc-50::lgc-50::GFP) rescued the
hyper-responsive phenotype of lgc-50 null animals to 30% 1-octanol in the absence of
food, indicating that the LGC-50::GFP fusion was localizing and functioning properly
(Figure 3.2.1). Confocal microscopy of lgc-50 null animals expressing the LGC-50::GFP
fusion protein (and stained with DiD to identity a subset of six sensory neurons to aid in
43
localization) indicated that lgc-50 was expressed in head muscle, as well as a number of
neurons, most notably the two ASI sensory neurons (Figure 3.2.5A-B). Surprisingly, in
contrast to reports on the PF4 receptor from parasitic nematodes, such as A. suum, only
very weak and variable expression was observed in body wall muscle (Figure 3.2.5C).
As predicted based on the expression data, RNAi knockdown and rescue of lgc-50 using
cell-selective promoters suggested that LGC-50 functioned in the ASIs (Pgpa-4 & Psra6) to modulate the aversive response to 30% 1-octanol (Figure 3.2.6 & 3.2.7). Although
LGC-50::GFP expression was not observed in the ASE or AWC sensory neurons (most
probably because of low expression levels), the hyper-responsive phenotype observed in
the absence of food and in the presence of elevated NaCl suggested that LGC-50 might
also be functioning in these neurons, as they have been implicated in salt/ volatile
odorants chemotaxis (Bargmann et al., 1993; Pierce-Shimomura et al., 2001; Bargmann,
2006; Suzuki et al., 2008). Indeed, RNAi knockdown and rescue using multiple
promoters also supports a role for lgc-50 in both the ASEs and AWCs (Figure 3.2.6 &
3.2.7). This phenomenon where RNAi knockdown in one pair of neurons is sufficient to
mimic the null phenotype, where expression in a different pair of neurons is sufficient to
rescue the null animals has been discussed previously and may result from overexpression during rescue (Mills et al., 2012). For example, ASI-selective (Pgpa-4) RNAi
knockdown of lgc-50 in wild type animals is sufficient to reproduce lgc-50 null hyperresponsiveness to 30% 1-octanol (5 s), while ASI-selective (Pgpa-4) rescue of lgc-50 in
lgc-50 null background can restore aversive response to wild type level (10 s) (Figure
3.2.6 & 3.2.7).
44
Figure 3.2.5. An Plgc-50::lgc-50(+)::GFP transgene is expressed in head muscle, as
well as a number of head and tail neurons, including the ASI sensory neurons and
the ventral cord motor neurons.
An lgc-50 translational transgene containing ~5kb upstream of the lgc-50 translational
start site (ATG) fused to GFP was injected into lgc-50 animals (A-C). Animals
expressing the Plgc-50::lgc-50(+)::GFP transgene (green) were incubated with DID (red)
to stain amphid and phasmid neurons for identification and co-localization (yellow).
Panel A: Anterior portion of a Plgc-50::lgc-50(+)::GFP expressing animal. Panel B: Inset
from Panel A of DID stained neurons. Panel C: Posterior portion of a Plgc-50::lgc50(+)::GFP expressing animal.
45
Figure 3.2.6. Aversive responses to 30% 1-octanol are more rapid after the neuronselective RNAi knockdown of lgc-50 in the ASI, ASH, ASE AWC and/or AWB
sensory neurons in wild type animals, mimicking the more rapid aversive phenotype
observed in lgc-50 null animals.
These neuron-selective RNAi results suggest LGC-50 may function at multiple locations
to inhibit aversive responses. Animals were examined on fresh NGM plates without
addition for their ability to respond to 30% 1-octanol, as we have described previously.
Data are presented as mean  SE and analyzed by a two-tailed Student‟s t test.
„*‟p<0.001, significantly different from wild-type animals assayed under identical
conditions.
46
Figure 3.2.7. The neuron-selective expression of lgc-50 in ASI, ASH, ASE AWC
and/or AWB sensory neurons in lgc-50 null animals can rescue the hyperresponsiveness (5 s) to 30% 1-octanol observed in lgc-50 null animals to wild type
level (10 s).
These neuron-selective rescues of lgc-50 further support the hypothesis that LGC-50 may
function at multiple locations to inhibit aversive responses. Animals were examined on
fresh NGM plates without addition for their ability to respond to 30% 1-octanol, as we
have described previously. Data are presented as mean  SE and analyzed by a two-tailed
Student‟s t test. „*‟p<0.001, significantly different from wild-type animals assayed under
identical conditions.
47
3.2.4. The direct injection of PF4 into animals over-expressing lgc-50 in
body wall muscle causes paralysis
In contrast to A. suum and other larger parasitic nematodes, lgc-50 does not appear to be
robustly expressed in C. elegans body wall muscle, although it is expressed in head
muscle and a number of neurons, based on the expression of Plgc-50::lgc-50::GFP
(Figure 3.2.5; Jex et al., unpublished). Not surprisingly, the direct injection of PF4 into
the pseudoecoelom of C. elegans did not cause paralysis, in contrast to its effects in A.
suum (Reinitz et al., 2011) (Figure 3.2.8). Therefore, to examine the relationship
between PF4 and LGC-50 more directly, we over-expressed lgc-50::GFP in body wall
muscle using the myo-3 promoter. As predicted, the direct injection of PF4 (100 μM)
into the pseudocoelom of animals over-expressing lgc-50 in the body wall muscle caused
a rapid onset of paralysis (Figure 3.2.8). This paralysis was temporary, as mobility was
restored about 20 min post-injection, and was not observed in wild type or lgc-50 null
animals, or sham-injected animals over-expressing lgc-50 in body wall muscle (data not
shown).
48
Figure 3.2.8. The direct injection of PF4 (100 µM) into the pseudocoelom of animals
over-expressing lgc-50 in the body wall muscle (Pmyo-3) causes rapid onset of
paralysis.
Mobility returned to the paralyzed animals at 20 min post injection. Animals were
examined on fresh NGM plates with no addition. Data are presented as mean  SE and
analyzed by a two-tailed Student‟s t test. „*‟p<0.001, significantly different from wildtype animals assayed under identical conditions. The number of animals examined is
indicated within each bar.
3.2.5. Animals over-expressing lgc-50 in body wall muscle have reduced
mobility in the presence of food
Food is reported to stimulate the release of flp-1 encoded peptides (Nelson et al.,
1998; Li et al. 1999). Animals over-expressing lgc-50 in body wall muscle move more
slowly on OP50 bacterial lawn than wild type animals (Figure 3.2.9A). This reduced
mobility is not observed when lgc-50 in over-expressed in the body wall muscle of flp-1
null animals. Furthermore, animals over-expressing lgc-50 in body wall muscle (Pmyo-3)
stop more frequently during forward locomotion in the presence of food, and again this
49
phenotype is not observed in when lgc-50 is over-expressed in the body all muscle of flp1 null animals (Figure 3.2.9B). Together, these two phenotypes provide further support
for the hypothesis that flp-1-encoded peptide(s) is a ligand for LGC-50.
Figure 3.2.9. Animals over-expressing lgc-50 in body wall muscle (Pmyo-3::lgc50::GFP transgene) move more slowly than wild-type animals in the presence of
food (OP50).
This sluggish phenotype is not observed in flp-1 animals over-expressing lgc-50 in body
wall muscle, providing yet another indicator that LGC-50 is the receptor for flp-1encoded peptides. Animals were examined on fresh NGM plates with food for their rate
of motility (body bends/ 20 s) (Fig. 16A) and frequency of pauses in 3 min (Fig. 16B).
Data are presented as mean  SE and analyzed by a two-tailed Student‟s t test.
„*‟p<0.001, significantly different from wild-type animals assayed under identical
conditions. The number of animals examined is indicated within each bar.
50
3.2.6. LGC-50 may form a heterologous channel with other ligandgated ion channel subunits
Repeated attempts to identify LGC-50 as a PF4 receptor after the heterologous
expression of lgc-50 in Xenopus oocytes have been unsuccessful, as no PF4-gated Clcurrents were observed in Xenopus oocytes expressing LGC-50. These negative results
could mean that 1) functional LGC-50 does not express well in oocytes, as has been
observed for other nematode receptors and ion channels (Bennett et al., 2012), 2) LGC50 is part of a heterologous channel and/or requires additional accessory proteins for
expression, as has been observed for other C. elegans ion channels (Boulin et al., 2008)
or 3) the LGC-50 is NOT the PF4 receptor. Indeed, the predicted protein sequence of
LGC-50 lacks a PARS domain (having instead, a unique SARS domain), a feature of the
pore-forming region (M2) of a typical Cl- channel, further suggesting that other
subunit(s) might be required to form a functional channel.
Assuming that LGC-50 requires other subunits to form a functional channel, we
predicted that null mutants for the other subunit(s) would also display phenotypes similar
to lgc-50 or flp-1 null animals. Indeed, a screen of 15 other potential ligand-gated ion
channels mutants using the octanol avoidance assay amazingly identified an additional
eight ion channel mutants that were hyper-responsive to 30% 1-octanol, lgc-34, lgc-40,
lgc-46, lgc-47, lgc-49, lgc-51, lgc-53 and lgc-54 (Figure 3.2.10). Further examination of
these mutants in the presence of TA revealed that lgc-34, lgc-46, lgc-47 and lgc-54
shared a similar phenotype to lgc-50 (Figure 3.2.11). This result was surprising and
suggests that ligand-gated Cl- channels provide a wealth of inhibitory modulatory inputs
into the locomotory circuit.
51
Figure 3.2.10. Many cys-loop, ligand-gated Cl- channel null mutants respond more
rapidly to 30% 1-octanol than wild-type animals.
Animals were examined on fresh NGM plates without addition for their ability to respond
to 30% 1-octanol, as we have described previously. Data are presented as mean  SE and
analyzed by a two-tailed Student‟s t test. „*‟p<0.001, significantly different from wildtype animals assayed under identical conditions.
52
Figure 3.2.11. LGC-34 and LGC-47 may form a heteromeric peptide-gated ion
channel with LGC-50.
Animals were examined on fresh NGM plates with 4 mM TA for their ability to respond
to 30% 1-octanol, as we have described previously. Data are presented as mean  SE and
analyzed by a two-tailed Student‟s t test. „*‟p<0.001, significantly different from wildtype animals assayed under identical conditions.
Interestingly, heat map transcriptome analysis in A. suum suggests that the A.
suum lgc-34 is very robustly expressed in body wall muscle throughout its life cycle (Jex
et al., unpublished). Similar expression patterns (in body wall muscles of adults and
larvae) have also been reported for the C. elegans lgc-34 (Meissner et al., 2011). Since
LGC-34 has clear orthologues in other parasitic nematodes, including Ascaris suum and
Dirofilaria immitis, we choose to examine lgc-34 null animals first (Yates &
Wolstenholme, 2004; Jex et al., unpublished). As predicted, if LGC-50 and LGC-34
form a heteromeric channel, lgc-34 over-expressors will also have aversive phenotypes
(to 30% 1-octanol) identical to lgc-50 over-expressors, i.e. both are stimulated by NaCl
53
and not inhibited by TA and OA (Figure 3.2.12). Furthermore, this hyper-responsiveness
to 30% 1-octanol was not present when lgc-50 was over-expressed in lgc-34 null
background, suggesting that LGC-50 and LGC-34 could form a heteromeric channel
gated by a flp-1-encoded peptide. These studies are continuing to characterize the
potential role of these additional cys-loop receptor subunits in a heteromeric PF4-gated
Cl- channel by expressing different combinations of these subunits (LGC-34 & LGC-47)
with LGC-50 in Xenopus oocytes.
Figure 3.2.12. Aversive responses in animals over-expressing lgc-50 or lgc-34 are
stimulated by NaCl and not inhibited by tyramine or octopamine.
In the presence of additional salt, both lgc-50 and lgc-34 over-expressors reverse more
rapidly in response to 30% 1-octanol than wild type animals, suggesting that LGC-50
might form a heteromeric channel with LGC-34. However, this enhanced aversive
response to 30% 1-octanol is not observed when lgc-50 is over-expressed in lgc-34
background, further indicating LGC-34 as a potential subunit partner with LGC-50 in a
heteromeric channel. Animals were examined on fresh NGM plates with additional 4 mM
NaCl, 4 mM TA or 4 mM OA for their ability to respond to 30% 1-octanol, as we have
54
described previously. Data are presented as mean  SE and analyzed by a two-tailed
Student‟s t test. „*‟p<0.001, significantly different from wild-type animals assayed under
identical conditions.
55
Chapter 4
Discussion
Most anthelmintics in use against infections of gut nematodes act as agonists at
key receptors in the neuromuscular junction and cause paralysis by interfering with
muscle contraction (locomotion), leading to parasite expulsion by peristalsis. As noted
above, agonists at four distinct neuromuscular molecular targets that interfere with
muscle contraction and cause paralysis have been successfully exploited as anthelmintics
against gut dwelling parasitic nematodes: two nicotinic ACh receptors
(tetrahydropyrimidines/imidathiazoles and amino-acetonitriles) and glutamate
(macrocyclic lactones)/ GABA (piperazine)-gated Cl- channels. (Martin, 1985; Geary et
al., 1993; Sheriff et al., 2002). However, except for the amino-acetonitrile derivatives,
resistance has begun to develop to all classes of anthelmintics, stressing the need for new
drug targets (Reynoldson et al., 1997; Bain, 1999; Albonico et al., 2002; Kaplan, 2004;
Wolstenholme et al., 2004). More importantly, successful anthelmintics against human
infections are still quite limited and have often been developed optimized first to combat
the veterinary parasites. For example, no useful, non-toxic treatment for adult filarial
infections, such as Wuchereria bancrofti, Brugia spp, Onchocerca volvulus, Loa loa is
currently available.
56
Monoamines, including 5-HT, TA, OA and DA, modulate most key behaviors in
nematodes, with monoaminergic signaling mediated by an array of G-protein coupled
receptors (GPCRs) and unique monoamine-gated Cl- channels. Exogenous 5-HT, DA or
TA also independently paralyze both free-living and parasitic nematodes, i.e., the
addition of monoamines can create uncoordinated, directionless movement, leading
ultimately to immobilized worms (Figure 3.1.1A & 3.1.2B). For example, exogenous 5HT causes an unusual “kinked” paralysis in J2s of the soybean cyst nematode Heterodera
glycines while 5-HT injection into A. suum causes immediate cessation of locomotory
waveforms (paralysis) (Reinitz & Stretton, 1996; Masler, 2007; Komuniecki et al., 2012).
This monoamine-mediated paralysis often appears distinct from the classical, spastic
paralysis initiated by cholinergic agonists, such as levamisole, or the flaccid paralysis
associated with GABA-ergic agonists that result from the activation of receptors directly
on body wall muscle. Indeed, paralysis in these monoamine-treated worms appears to
result most often from the disruption of complex locomotory decision-making networks
and the worms appear to be as much confused as paralyzed (Pokala et al., 2014; Law et
al., 2015; Summers et al., 2015). However, the identification of new targets within key
signaling pathways has been limited by the lack of useful information about the identity,
function and localization of the additional receptors regulating muscle contraction and
locomotion. Furthermore, we also need new high-throughput screening protocols that
preserve the unique pharmacologies of the receptors from the different parasites and
maintain a nematode-specific context that includes the cuticle and appropriate accessory
proteins, especially given that no nematode cells lines are available and the parasites
themselves are extremely difficult and expensive to culture.
57
C. elegans has been used in the past for large-scale small molecule screens and
chemical genomics and predictive models for drug accumulation and bioactivity have
been developed that may be used to bias preliminary screening (Burns et al., 2006; Burns
et al., 2010). These studies expand these previous observations and validate the use of
“chimeric” mutant C. elegans, created by the heterologous, ectopic expression of key
drug targets from parasitic nematodes, for use as a primary platform for anthelmintic
screening, target identification and potentially receptor deorphanization. This screening
approach is especially useful because nematode-specific cells lines are not available and
the expression of nematode receptors in mammalian cells is quite variable and can
require a host of additional modifications, including temperature shock to achieve
expression (Kubiak et al., 2003b; Kubiak et al., 2008; Larsen et al., 2012). In fact, few
studies have compared receptor pharmacologies in vivo with those of the nematode
receptors heterologously expressed in mammalian cells. In addition, this screening
platform also includes the nematode cuticle, a potential barrier to the entry of any
anthelmintic, as well as a wide array of ABC transporters involved in drug efflux and
resistance and, most importantly, appears to maintain the individual pharmacologies of
receptors from different parasitic nematodes, while providing the environment and
accessory proteins necessary for functional expression (Ardelli, 2013). As noted above,
although C. elegans cuticle appears to be more impermeable than those of some parasitic
nematodes, the permeability of the C. elegans cuticle can be manipulated by modifying
incubation conditions and availability of various mutant backgrounds (Chase et al., 2004;
Partridge et al., 2008; Schultz et al., 2014; Law et al., 2015). Although the present study
has focused on inhibitory GPCRs and ligand-gated ion channels, this screening approach
58
can potentially be expanded to any signaling molecules for which the appropriate mutant
backgrounds can be prepared. Specific promoters are available for C. elegans muscles
and most neurons; alternatively, specific promoters to other neurons can be generated
using a Cre-Lox approach (Zhang et al., 2008).
C. elegans has often been demeaned as a useful platform for anthelmintic
discovery in the past, mainly based on the preconceptions that signaling pathways will
differ dramatically between free-living and parasitic nematodes and that the free-living C.
elegans has cuticle that is highly impermeable to exogenously applied chemicals,
compared to parasitic nematodes, necessitating large amounts of potentially rare and
costly compounds for testing. Indeed, free-living nematodes are potentially exposed to
more environmental toxins than parasitic nematodes that reside in the relatively safe,
temperature-controlled, prescreened vertebrate gut. However, as described more fully
below, research in the past years has highlighted the conservation of the core signaling
pathways involved in locomotory decision-making with both large and small, free-living
and parasitic nematodes, having the same ~300 neurons with highly conserved molecular
characteristics (Brownlee et al., 1994; Johnson et al., 1996; Kim & Li, 2004; Johnston et
al., 2010; Rao et al., 2011; McCoy et al., 2014). In contrast, cuticular permeability has
been a potential problem for anthelmintic screening. For example, the recognized
anthelmintic derquantel (2-desoxoparahequamide) appears to function as a nicotinic
receptor antagonist and has marked activity in dissected C. elegans, but not intact worms,
suggesting that the permeability of the C. elegans cuticle may be more limited than that
of the parasites (Ruiz-Lancheros et al., 2011). The C. elegans cuticle is made up of six
layers, the epicuticle, external cortical, internal cortical, medial, fiber and basal, as well
59
as a carbohydrate-rich surface coat external to the epicuticle (Riddle et al., 1997a; Riddle
et al., 1997b). The lipid-rich epicuticle layer might be the key barrier to externallyapplied drugs, especially water-soluble molecules (5-HT, TA, 8OH-DPAT etc.) and the
reason for the high concentrations required to cause paralysis in isotonic environments,
i.e. on NGM agar plates (Riddle et al., 1997a; Riddle et al., 1997b; Page & Johnstone,
2007). However, as described above, the permeability of the C. elegans cuticle can be
manipulated by modifying incubation conditions and the use of mutants with altered
cuticle permeability. By incubating the animals in a salt-free, hypotonic environment, 5HT paralyzes wild-type animals with an EC50 of about 0.5 mM, in contrast with an EC50
of about 12 mM on isotonic NGM agar plates (Figure 3.1.1A-D). Similar increases in
permeability have also been observed when incubating wild type C. elegans in diluted
M9 buffer (1:1 in water) containing 1 mM 5-HT; 1 mM 5-HT has no effect on wild type
animals in M9 buffer alone (Komuniecki & Law, unpublished). In addition, a number of
C. elegans mutants that appear to have increased cuticular permeabilities may also be
useful for enhancing small molecule screening against an array of medically-important
targets, including those involved in locomotory paralysis (Partridge et al., 2008; Schultz
et al., 2014). For example, many of the bus (bacterially swollen) mutations appear to
alter the cuticle and increase permeability, even under isotonic conditions; indeed, the
hypotonic assay conditions uses in the present study cause uncontrolled swelling and are
lethal to the bus mutants examined (Figure 3.1.1E-F) (Partridge et al., 2008).
Additionally, as shown in Figure 3.1.1E-F, it might be possible to select specific cuticle
mutants with permeabilities that mimic those of individual parasitic nematodes, providing
a means to bypass the complicated and expensive process of culturing live parasites, at
60
least during preliminary stages of agonist screening. This ability to alter cuticular
permeability will certainly be useful for agonist and potential anthelmintic identification,
but in the case of the monoamines examined, relatively high concentrations of ligand are
still required and, ultimately, any potential agonists identified using this approach will
have to be validated in the intended target parasites.
As noted above, although nematodes vary tremendously in size (about ~1 mm for
C. elegans compared with ~300 mm for Ascaris suum), their body plans are remarkably
conserved, with adults of both species exhibiting nearly identical neuronal wiring
diagrams, as well as core signaling pathways and the locomotory machinery (Brownlee et
al., 1994; Johnson et al., 1996; Jarecki et al., 2010). For example, both A. suum and C.
elegans have a nerve ring around the pharynx and similar number of neurons at the adult
stage, even though an adult hermaphrodite C. elegans has only 959 cells, compared to
~10000 muscle cells alone in an adult Ascaris (Stretton, 1976). Furthermore, all
commercially available anthelmintics appear to have similar activity against C. elegans as
parasitic nematodes, and our understanding of their modes of action has, in large part,
resulted from our ability to genetically manipulate their putative targets in respective C.
elegans mutant backgrounds (Holden-Dye et al., 2012; Krucken et al., 2012; Miltsch et
al., 2013; Hernando & Bouzat, 2014). These observations suggest that C. elegans, with
its well-defined molecular genetics, numerous signaling mutants and cell-based assay
systems might be a useful model to identify core signaling pathways in parasitic
nematodes and could provide unique new insights, compared with studies focused
exclusively in parasites. For example, the current study has identified signaling in the
two AIB interneurons as a potential target for anthelmintic development, since the
61
inhibition of AIB signaling via activation of ligand-gated Cl- channel or Gαo-coupled
GPCR causes locomotory confusion and subsequent paralyze (Figure 3.1.2b & 3.1.7).
Indeed it would be reasonable to assume that any receptor that reduces AIBs signaling
(and consequently, signaling in corresponding downstream interneurons) can be exploited
as a drug target. Furthermore, the use of the more extensively studied and experimentally
pliable C. elegans will expedite the rapid and precise identification of potential secondary
targets for the current anthelmintics, as demonstrated in the discovery in the present study
of DOP-3 as another major target for PAPP, apart from the previously identified 5-HT1like receptor (Figure 3.1.4). It would be interesting to determine if the in vivo activity of
PAPP in clearing H. contortus infections was due to its agonism of 5-HT or DA
signaling, or if both pathways were required (Smith et al., 2003; White et al., 2007).
These observations suggest that our screening protocols might also be useful in
identifying and enhancing the activity of additional anthelmintics on potential secondary
targets should resistance (mutation) arise in the primary targets.
The recent availability and expanded analysis of many nematode genomes has
supported the hypothesis that many core signaling pathways are highly conserved in both
free-living and parasitic nematodes (Geary & Thompson, 2001; Gilleard et al., 2005;
Brown et al., 2006; Ghedin et al., 2007; Jex et al., 2011; Laing et al., 2013; Schwarz et
al., 2013). Additionally, most, if not all C. elegans receptors have their respective
orthologous counterparts in parasitic nematodes of significant importance. For example,
15 of the 16 C. elegans monoamine receptors have clear orthologues in the recently
completed A. suum genome, even though these animals diverged over hundreds of
millions of years ago (Jex et al., 2011) (Figure 1.1.1 & 1.1.2). However, nematodes do
62
exhibit significant diversity, so that there is no guarantee that processes in C. elegans will
be exactly duplicated in parasitic nematodes. Indeed, physiological, biochemical and/or
molecular differences between or among nematode species have been demonstrated. For
example, the composition of gene families and individual splicing patterns can vary
significantly within the phylum. Conspicuous examples include tyra-2 and tyra-3, both
of which are represented by three isoforms in C. elegans, with distinct homologues
detected for each isoform in A. suum (Jex et al., 2011). Another notable difference is the
time (developmental stage) of expression and site of action. For example, the 5-HTgated Cl- channel, MOD-1 appears to be robustly expressed in body wall muscle during
larval stages of C. elegans development, but more limited to neurons (including AIBs) in
adult (Gurel et al., 2012). Additionally, due to the drastic differences in “lifestyle”,
orthologous receptors may not be working at the same anatomical locations in C. elegans
and its parasitic cousins, as in the case of the putative FLP-1/PF4-gated Cl- channel
subunits, LGC-50 & LGC-34, which is reported to be robustly expressed in the body wall
muscle of A. suum (mRNA heat map & electrophysiology data), but is primarily localized
to the head muscles and neurons in the nerve rings in C. elegans (GFP-tagged
translational fusion) (Figure 3.2.5) (Maule et al., 1995a; Maule et al., 1995b; Jex et al.,
unpublished). However, recent work suggests that GFP expression can be promiscuous
or, alternatively, some genes are functionally expressed in neurons not exhibiting GFP
fluorescence (from low expression level) (Ezak & Ferkey, 2010; Ezcurra et al., 2011).
These caveats emphasize that any observations from C. elegans need to be confirmed by
direct assay in individual parasite species wherever possible.
63
We, as well as other researchers, have demonstrated the rescue of C. elegans null
mutants with their respective orthologues from parasites (Crisford et al., 2011; Welz et
al., 2011; Komuniecki & Law, unpublished). For example, ivermectin and emodepside
paralyze both C. elegans and parasitic nematodes through the activation of orthologous
glutamate-gated Cl- and SLO-1 channels, respectively, and the expression of these
receptors from parasitic nematodes can rescue the appropriate C. elegans null mutants,
further validating the utility of our “dual systems” approach for target identification
(Glendinning et al., 2011; Welz et al., 2011). Indeed, the current study has highlighted
C. elegans as a promiscuous expression platform, not only of proteins from other
parasitic nematodes (H. contortus, in current study), but also those from origin as diverse
as mammal (human) and insect (Drosophila). More importantly, these proteins are
functional and appear to retain their ligand selectivity, implying that chimeric C. elegans
can be used to screen for a nematode-selective compounds with minimal cross-activity to
the intended host receptors, as shown in Figure 3.1.3B where only 5-HT quint animals
expressing the human 5-HT1A receptor (HTR1A) are paralyzed by the subtype-selective
5-HT1A receptor agonist, 8-OH-DPAT, while animals with either nematode (SER-4) or
Drosophila (5HT1A) are not affected. Interestingly, in addition to the human 5-HT1
receptor, we have also demonstrated the robust expression of human kappa opioid and
cannabinoid receptors in C. elegans (Komuniecki, Mills & Oakes, unpublished). It is
unclear why human receptors can be expressed so effectively in C. elegans, when the
converse is not usually true, but one reason may be that the GIRK/ arrestin system
responsible for GPCR down regulation in human functions quite differently in C.
64
elegans, meaning that once the human receptor are activated they do not desensitize in a
C. elegans background (Pearce et al., 2010; Gurevich et al., 2012).
The challenge of using a non-nematode heterologous expression system such as
Xenopus oocytes or other cell-based systems has been highlighted in many previous
studies, including our ongoing attempts to functionally express the putative FLP-1/PF4gated Cl- channel subunits, LGC-50 and LGC-34 in Xenopus oocytes (Kubiak et al.,
2003b; Kubiak et al., 2008; Larsen et al., 2012). For a heteromeric cys-loop ligand-gated
ion channel, many accessories proteins in addition to the channel subunits themselves (up
to five different subunits) are often required for functional expression (Boulin et al.,
2008; Boulin et al., 2011; Bennett et al., 2012). Additionally, changes in the ratio of
subunits can result in significant changes in ligand sensitivity/selectivity. For example,
changing the ratio of two A. suum nicotinic ACh receptor subunits, UNC-29 & UNC-38
causes the formation of distinct receptors with different ligand selectivity/ sensitivity
(Williamson et al., 2009). Certainly, LGC-34/LGC-50 alone may not be sufficient to
express a functional FLP-1/PF4 receptor. However, substantial and robust geneticbehavioral data support the hypothesis that LGC-34 and LGC-50 are essential for the
formation of a Cl- channel gated by PF4 or other FLP-1-encoded neuropeptide(s). For
example 1) flp-1, lgc-50 and lgc-34 exhibit similar null and over-expression phenotypes,
2) lgc-34 & lgc-50 over-expression phenotypes are not observed in flp-1 null
backgrounds and 3) lgc-50 over-expression phenotypes are absent in an lgc-34 null
background, suggesting that either LGC-50 forms a common receptor with LGC-34, or
LGC-50 and LGC-34 are part of one signaling pathway, with LGC-50 potentially located
upstream of LGC-34 (Figure 3.2.12). Perhaps the most direct indicator of LGC-50
65
involvement in PF4-mediated paralysis is that the direct injection of PF4 into the
pseudocoelom of wild type animals over-expressing LGC-50 in body wall muscle (Pmyo3) caused paralysis, in contrast to PF4 injection into either non-transgenic wild type
animals or lgc-50 null mutants (Figure 3.2.8). The observation that non-transgenic wildtype animals were not paralyzed by PF4 injection supports the previous observation that
LGC-50 is not robustly expressed in body wall muscle (Figure 3.2.5). The transient
nature of paralysis suggests the rapid degradation of injected PF4 or receptor
desensitization. Further screening will be required to identify other potential PF4
receptor subunit(s), most likely by examining additional receptor combinations or
different heterologous expression systems. These attempts to characterize LGC-50/LGC34 highlighted the utility of expression in transgenic C. elegans to initially bypass the
multitudes of obstacles mentioned previously for other heterologous expression systems.
In summary, the present study has identified and validated a novel approach to
anthelmintic screening, using chimeric mutants C. elegans expressing key drug targets
from parasitic nematodes at sites yielding robust locomotory phenotypes upon agonist
stimulation. Using this approach, we have identified 1) selective agonists for a nematode
5-HT1-like receptor, as key target of 5-HT dependent paralysis, 2) a key role for the two
AIB interneurons in mediating “locomotory confusion” and paralysis, 3) an additional
target for PAPP, a well characterized anthelmintic capable of clearing H. contortus
infections from gerbils, and 4) cys-loop receptor subunits involved in the flaccid paralysis
associated with the widely-expressed nematode neuropeptide, PF4. These studies are
continuing to identify additional 5-HT1-like agonists as potential lead compounds for
66
anthelmintic development and to firmly establish the subunit identity of the PF4 receptor
in parasitic nematodes.
67
References
Ailion M & Thomas JH. All available mutations in flp-1 also delete daf-10 sequences,
confounding phenotype interpretation. Worm Breeder's Gazette 2001; 16(5): 41
Albonico M, Bickle Q, Haji HJ, Ramsan M, Katrib KJ, Montresor A, Savioli L & Taylor
M. Evaluation of the efficacy of pyrantel-oxantel for the treatment of soil-transmitted
nematode infections. Trans R Soc Trop Med Hyg 2002; 96(6): 685-90
Allen AT, Maher KN, Wani KA, et al. Coexpressed D1- and D2-like dopamine receptors
antagonistically modulate acetylcholine release in Caenorhabditis elegans. Genetics
2011; 188: 579-590
Ardelli BF. Transport proteins of the ABC systems superfamily and their role in drug
action and resistance in nematodes. Parasitol Int 2013; 62(6): 639-46
Awasthi S & Bundy D. Intestinal nematode infection and anaemia in developing
countries. BMJ 2007; 334: 1065-66
68
Bain RK. Irradiated vaccines for helminth control in livestock. Int J Parasitol 1999; 29:
185-191
Bargmann CI, Hartwieg E & Horvitz R. Odorant-selective genes and neurons mediate
olfaction in C. elegans. Cell 1993; 74(3): 515-27
Bargmann CI. Chemosensation in C. elegans. WormBook 2006; 1-29
Beech RN, Callanan MK, Rao VT et al. Characterization of cys-loop receptor genes
involved in inhibitory amine neurotransmission in parasitic and free living nematodes.
Parasitol Int 2013; 62: 599-605
Bennett HM, Lees K, Harper KM, Jones AK, Sattelle DB, Wonnacott S & Wolstenholme
AJ. Xenopus laevis RIC-3 enhances the functional expression of the C. elegans
homomeric nicotinic receptor, ACR-16, in Xenopus oocytes. J Neurochem 2012; doi:
10.1111/jnc.12013
Boulin T, Gielen M, Richmond JE, Williams DC, Paoletti P & Bessereau JL. Eight genes
are required for functional reconstitution of the Caenorhabditis elegans levamisolesensitive acetylcholine receptor. Proc Natl Acad Sci U S A 2008; 105(47): 18590-5
Boulin T, Fauvin A, Charvet CL, Cortet J, Cabaret J, Bessereau JL & Neveu C.
Functional reconstitution of Haemonchus contortus acetylcholine receptors in Xenopus
69
oocytes provides mechanistic insights into levamisole resistance. Br J Pharmacol 2011;
164(5): 1421-32
Brooker S. Estimating the global distribution and disease burden of intestinal nematode
infections: adding up the numbers - a review. Int J Parasitol 2010; 40: 1137-1144
Brown LA, Jones AK, Buckingham SD, Mee CJ & Sattelle DB. Contributions from
Caenorhabditis elegans functional genetics to antiparasitic drug target identification and
validation: nicotinic acetylcholine receptors, a case study. Int J Parasitol 2006; 36: 617-24
Brownlee DJ, Fairweather I, Johnston CF & Shaw C. Immunocytochemical
demonstration of peptidergic and serotoninergic components in the enteric nervous
system of the roundworm, Ascaris suum (Nematoda, Ascaroidea). Parasitology 1994;
108: 89-103
Buckingham SD & Sattelle DB. Fast, automated measurement of nematode swimming
(thrashing) without morphometry. BMC Neurosci 2009; 10: 84
Burns AR, Kwok TC, Howard A, et al. High-throughput screening of small molecules for
bioactivity and target identification in Caenorhabditis elegans. Nat Protoc 2006; 1: 190614
70
Burns AR, Wallace IM, Wildenhain J, et al. A predictive model for drug bioaccumulation
and bioactivity in Caenorhabditis elegans. Nat Chem Biol 2010; 6: 549-57
Carr JA, Parashar A, Gibson R, et al. A microfluidic platform for high-sensitivity, realtime drug screening on C. elegans and parasitic nematodes. Lab Chip 2011; 11: 2385-96
Carswell CL, Sun J & Baenziger JE. Intramembrane aromatic interactions influence the
lipid sensitivities of pentameric ligand-gated ion channels. J Biol Chem. 2015; 290(4):
2496-507
Chao MY, Komatsu H, Fukuto HS, Dionne HM & Hart AC. Feeding status and serotonin
rapidly and reversibly modulate a Caenorhabditis elegans chemosensory circuit. Proc
Natl Acad Sci U S A. 2004; 101(43): 15512-7
Chase DL, Pepper JS & Koelle MR. Mechanism of extrasynaptic dopamine signaling in
Caenorhabditis elegans. Nat Neurosci. 2004; 7: 1096-1103
Chen B, Deutmeyer A, Carr J, et al. Microfluidic bioassay to characterize parasitic
nematode phenotype and anthelmintic resistance. Parasitology 2011; 138: 80-8
Cowden C & Stretton AOW. Eight novel FMRFamide-like neuropeptides isolated from
the nematode Ascaris suum. Peptides 1995; 16(3): 491-500
71
Crisford A, Murray C, O‟Connor V, et al. Selective toxicity of the anthelmintic
emodepside revealed by heterologous expression of human KCNMA1 in Caenorhabditis
elegans. Mol Pharmacol 2011; 79: 1031-1043
Donnelly JL, Clark CM, Leifer AM, et al. Monoaminergic orchestration of motor
programs in a complex C. elegans behavior. PLoS Biol 2013; 11: e1001529
Durrnagel S, Kuhn A, Tsiairis CD, Williamson M, et al. Three homologous subunits
form a high affinity peptide-gated ion channel in Hydra. J Biol Chem 2010; 285(16):
11958-65
Ezak MJ & Ferkey DM. The C. elegans D2-like dopamine receptor DOP-3 decreases
behavioral sensitivity to the olfactory stimulus 1-octanol. PLoS One 2010; 5(2): e9487.
Ezcurra M, Tanizawa Y, Swoboda P & Schafer WR. Food sensitizes C. elegans
avoidance behaviors through acute dopamine signaling. EMBO J 2011; 30: 1110-22
Furukawa Y, Miyawaki Y & Abe G. Molecular cloning and functional characterization of
the Aplysia FMRFamide-gated Na+ channel. Pflugers Arch 2006; 451(5): 646-56
Geary TG, Sims SM, Thomas EM, et al. Haemonchus contortus: ivermectin-induced
paralysis of the pharynx. Exp Parasitol 1993; 77: 88-96
72
Geary TG, Marks NJ, Maule AG, Bowman JW, Alexander-Bowman SJ, Day TA, Larsen
MJ, Kubaik TM, Davis JP & Thompson DP. Pharmacology of FMRFamide-related
peptides in helminths. Ann N Y Acad Sci 1999; 897: 212-27
Geary TG & Thompson DP. Caenorhabditis elegans: how good a model for veterinary
parasites? Vet Parasitol 2001; 101: 371-86
Geary TG. Ivermectin 20 years on: maturation of a wonder drug. Trends Parasitol 2005;
21: 530-532
Geary TG, Woo K, McCarthy JS, Mackenzie CD, Horton J, Prichard RK, de Silva NR,
Olliaro PL, Lazdins-Helds JK, Engels DA & Bundy DA. Unresolved issues in
anthelmintic pharmacology for helminthiases of humans. Int J Parasitol 2010; 40: 1-13
Ghedin E, Wang S, Spiro D, Caler E, Zhao Q, Crabtree J, Allen JE, Delcher AL, Guiliano
DB, Miranda-Saavedra D et al. Draft genome of the filarial nematode parasite Brugia
malayi. Science 2007; 317(5845): 1756-60
Gill JH, Kerr CA, Shoop WL & Lacey E. Evidence of multiple mechanisms of
avermectin resistance in Haemonchus contortus - comparison of selection protocols. Int J
Parasitol 1998; 28: 783-9
73
Gilleard JS, Woods DJ & Dow JA. Model-organism genomics in veterinary parasite
drug-discovery. Trends Parasitol 2005; 21: 302-5
Glendinning SK, Buckingham SD, Sattelle DB, Wonnacott S & Wolstenholme AJ.
Glutamate-gated chloride channels of Haemonchus contortus restore drug sensitivity to
ivermectin-resistant Caenorhabditis elegans. PLoS One 2011; 6: e22390
Gray JM, Hill JJ & Bargmann CI. A circuit for navigation in Caenorhabditis elegans.
Proc Natl Acad Sci U S A 2005; 102(9): 3184-91
Gurel G, Gustafson MA. Pepper JS, et al. Receptors and other signaling proteins required
for serotonin control of locomotion in Caenorhabditis elegans. Genetics 2012; 192:
1359-71
Gurevich, EV, Tesmer, JJ, Mushegian, A & Gurevich, VV (2012) G protein-coupled
receptor kinases: more than just kinases and not only for GPCRs. Pharmacol Ther 2012;
133: 40-69
Hapiak V, Hobson R, Hughes L, et al. Dual excitatory and inhibitory serotonergic inputs
modulate egg-laying in Caenorhabditis elegans. Genetics 2009; 181: 153-163
74
Hapiak V, Summers P, Ortega A, Law WJ et al. Neuropeptides amplify and focus the
monoaminergic inhibition of nociception in Caenorhabditis elegans. J Neurosci 2013;
33(35): 14107-16
Harris G, Hapiak V, Wragg R, Miller S, Smith K, Hughes L, et al. Three distinct amine
receptors operating a different levels within the locomotory circuit are each essential for
the serotonergic modulation of chemosensation in Caenorhabditis elegans. J Neurosci
2009; 29: 1446-56
Harris G, Mills H, Wragg R, Hapiak V, et al. The monoaminergic modulation of sensorymediated aversive responses in Caenorhabditis elegans requires glutamatergic/
peptidergic cotransmission. J Neurosci 2010; 30(23): 7889-99
Harris G, Korchnak A, Summers P, Hapiak V, et al. Dissecting the Serotonergic Food
Signal Stimulating Sensory-Mediated Aversive Behavior in C. elegans. PLoS One 2011;
6(7): e21897
Hernando G & Bouzat C. Caenorhabditis elegans neuromuscular junction: GABA
receptors and ivermectin action. PLoS One 2014; 9: e95072
Ho NF, Geary TG, Barsuhn CL, et al. Mechanistic studies in the transcuticular delivery
of antiparasitic drugs. II: Ex vivo/in vitro correlation of solute transport by Ascaris suum.
Mol Biochem Parasitol 1992; 52: 1-13
75
Hobert O. PCR fusion-based approach to create reporter gene constructs for expression
analysis in transgenic C. elegans. Biotechniques 2002; 32: 728-30
Hobson RJ, Hapiak VM, Xiao H, Buehrer KL & Komuniecki R. SER-7: a
Caenorhabditis elegans 5-HT7-like receptor is essential for the 5-HT stimulation of
pharyngeal pumping and egg-laying. Genetics 2006; 172: 159-169
Holden-Dye L, Brownlee DJA & Walker RJ. The effects of the peptide KPNFIRFamide
(PF4) on the somatic muscle cells of parasitic nematode Ascaris suum. Br J Pharmacol
1997; 120(3): 379-386
Holden-Dye L, Crisford A, Welz C, et al. Worms take to the slo lane: a perspective on
the mode of action of emodepside. Invert Neurosci 2012; 12: 29-36
Hotez PJ, Brindley PJ, Bethony JM, King CH, Pearce EJ & Jacobson J. Helminth
infections: the great neglected tropical diseases. J Clin Invest 2008; 118(4): 1311-21
Hotez PJ & Kamath A. Neglected tropical diseases in sub-Saharan Africa: review of their
prevalence, distribution and disease burden. PLoS Negl Trop Dis 2009; 3(8): e412
76
Jarecki JL, Andersen K, Konop CJ, et al. Mapping neuropeptide expression by mass
spectrometry in single dissected identified neurons from the dorsal ganglion of the
nematode Ascaris suum. ACS Chem Neurosci 2010; 1: 505-19
Jex AR, Liu S, Li B, Young ND, Hall RS, Li Y, Yang L, Zeng N, Xu X, Xiong Z et al.
Ascaris suum draft genome. Nature 2011; 479(7374): 529-33
Johnson CD, Reinitz CA, Sithigorngul P & Stretton AO. Neuronal localization of
serotonin in the nematode Ascaris suum. J Comp Neurol 1996; 367: 352-60
Johnston MJ, McVeigh P, McMaster S, et al. FMRFamide-like peptides in root knot
nematodes and their potential role in nematode physiology. J Helminthol 2010; 84(3):
253-65
Jones AK & Sattelle DB. The cys-loop ligand-gated ion channel gene superfamily of the
nematode, Caenorhabditis elegans. Invert Neurosci 2008; 8(1): 41-7
Jones JT, Haegeman A, Danchin EGJ, et al. Top 10 plant-parasitic nematodes in
molecular plant pathology. Mol Plant Pathol 2013; 14: 946-961
Kaminsky R, Ducray P, Jung M, Clover R, Rufener L, Bouvier J, et al. A new class of
anthelmintics effective against drug-resistant nematodes. Nature 2008; 452: 176-180
77
Kaplan RM. Drug resistance in nematodes of veterinary importance: a status report.
Trends Parasitol 2004; 20: 477-481
Keating J, Yukich JO, Mollenkopf S & Tediosi F. Lymphatic filariasis and
onchocerciasis prevention, treatment and control costs across diverse settings: a
systematic review. Acta Trop 2014; 135C: 86-95
Kim K & Li C. Expression and regulation of an FMRFamide-related neuropeptide gene
family in Caenorhabditis elegans. J Comp Neurol 2004; 475(4): 540-50
Komuniecki R, Law W, Jex A, et al. Monoaminergic signaling as a target for
anthelmintic drug discovery: receptor conservation among the free-living and parasitic
nematodes. Mol Biochem Parasitol 2012; 183: 1-7
Krucken J, Harder A, Jeschke P, et al. Anthelmintic cyclcooctadepsipeptides: complex in
structure and mode of action. Trends Parasitol 2012; 28: 385-94
Kubiak TM, Larsen MJ, Davis JP, Zantello MR & Bowman JW. AF2 interaction with
Ascaris suum body wall muscle membranes involves G-protein activation. Biochem
Biophys Res Commun 2003a; 301(2): 456-9
78
Kubiak TM, Larsen MJ, Zantello MR, et al. Functional annotation of the putative orphan
Caenorhabditis elegans G-protein-coupled receptor C10C6.2 as a FLP15 peptide
receptor. J Biol Chem 2003b; 278: 42115-20
Kubiak TM, Larsen MJ, Bowman JW, et al. FMRFamide-like peptides encoded on the
flp-18 precursor gene activate two isoforms of the orphan Caenorhabditis elegans Gprotein-coupled receptor Y58G8A.4 heterologously expressed in mammalian cells.
Biopolymers 2008; 90: 339-48
Laing R, Kikuchi T, Martinelli A, Tsai IJ, Beech RN, Redman E, Holroyd N, Bartley DJ,
Beasley H, Britton C et al. The genome and transcriptome of Haemonchus contortus, a
key model parasite for drug and vaccine discovery. Genome Biol 2013; 14(8): R88
Larsen MJ, Lancheros ER, Williams T, et al. Functional expression and characterization
of the C. elegans G-protein-coupled FLP-2 Receptor (T19F4.1) in mammalian cells and
yeast. Int J Parasitol Drugs Drug Resist 2012; 15: 1-7
Law W, Wuescher LM, Ortega A, Hapiak VM, et al. Heterologous Expression in
Remodeled C. elegans: A Platform for Monoaminergic Agonist Identification and
Anthelmintic Screening. PLoS Pathog. 2015; 11(4): e1004794
Li C, Nelson LS, Kim K, Nathoo A & Hart AC. Neuropeptide gene families in the
nematode Caenorhabditis elegans. Ann N Y Acad Sci 1999; 897: 239-252
79
Li C & Kim K. Neuropeptide gene families in Caenorhabditis elegans. Adv Exp Med
Biol 2010; 692: 98-137
Li C. The ever-expanding neuropeptide gene families in the nematode Caenorhabditis
elegans. Parasitology 2005; 131
Lingueglia E, Deval E & Lazdunski M. FMRFamide-gated sodium channel and ASIC
channels: a new class of ionotropic receptors for FMRFamide and related peptides.
Peptides 2006; 27(5): 1138-52
Lustigman S & McCarter JP. Ivermectin resistance in Onchocerca volvulus: toward a
genetic basis. PLoS Negl Trop Dis 2007; 1(1): e76
Martin RJ. gamma-Aminobutyric acid- and piperazine-activated single-channel currents
from Ascaris suum body muscle. Br J Pharmacol 1985; 84: 445-61
Martin RJ & Robertson AP. Control of nematode parasites with agents acting on
neuromusculature systems: Lessons for neuropeptide ligand discovery. Adv Exp Med
Biol. 2010; 692: 138-154
Martin RJ, Buxton SK, Neveu C, et al. Emodepside and SLO-1 potassium channels: a
review. Exp Parasitol 2012; 132: 40-6
80
Masler EP. Responses of Heterodera glycines and Meloidogyne incognitato exogenously
applied neuromodulators. J Helminthol 2007; 81: 421-427
Maule AG, Shaw C, Bowman JW, Halton DW, Thompson DP, Thim L et al. Isolation
and preliminary biological characterization of KPNFIRFamide, a novel FMRFamiderelated peptide from the free-living nematode, Panagrellus redivivus. Peptides 1995a;
16(1): 87-93
Maule AG, Geary TG, Bowman JW, Marks NJ, Blair KL, Halton DW et al. Inhibitory
effects of nematode FMRFamide-related peptides (FaRPs) on muscle strips from Ascaris
suum. Invert Neurosci 1995b; 1(3): 255-65
Maule AG, Geary TG, Bowman JW, Shaw C, Halton DW & Thompson DP. The
Pharmacology of nematode FMRFamide-related peptides. Parasitol Today 1996; 12(9):
351-57
McCoy CJ, Atkinson LE, Zamanian M, et al. New insights into the FLPergic
complements of parasitic nematodes: Informing deorphanisation approaches. EuPA Open
Proteom 2014; 3: 262-272
81
McDonald PW, Hardie SL, Jessen TN, et al. Vigorous motor activity in Caenorhabditis
elegans requires efficient clearance of dopamine mediated by synaptic localization of the
dopamine transporter DAT-1. J Neurosci 2007; 27: 14216-27
McVeigh P, Leech S, Mair GR, Marks NJ, Geary TG & Maule AG. Analysis of
FMRFamide-like peptide (FLP) diversity in phylum Nematoda. Int J Parasitol 2005;
35(10): 1043-60
McVeigh P, Geary TG, Marks NJ & Maule AG. The FLP-side of nematodes. Trends
Parasitol 2006; 22(8): 385-96
Meissner B, Rogalski T, Viveiros R, Warner A, Plastino L, Lorch A, Granger L, Segalat
L & Moerman DG. Determining the sub-cellular localization of proteins within
Caenorhabditis elegans body wall muscle. PLoS One 2011; 6(5): e19937
Mello C & Fire A. DNA Transformation. Methods Cell Biol 1995; 48: 451-82
Mills H, Wragg R, Hapiak V, Castelletto M, et al. Monoamines and neuropeptides
interact to inhibit aversive behaviour in Caenorhabditis elegans. EMBO J 2012;
31(3):667-78
82
Miltsch SM, Krucken J, Demeler J, et al. Interactions of anthelmintic drugs in
Caenorhabditis elegans neuro-muscular ion channel mutants. Parasitol Int 2013; 62(6):
591-8
Nelson LS, Rosoff ML & Li C. Disruption of a neuropeptide gene, flp-1, causes multiple
behavioral defects in Caenorhabditis elegans. Science 1998; 281(5383): 1686-90
Page AP & Johnstone IL. The cuticle. In: WormBook: The online review of C. elegans
biology 2007
Available: http://www.ncbi.nlm.nih.gov/books/NBK19745/
Accessed: 10 January 2015
Partridge FA, Tearle AW, Gravato-Nobre MJ, Schafer WR & Hodgkin J. The C. elegans
glycosyltransferase BUS-8 has two distinct and essential roles in epidermal
morphogenesis. Dev Biol 2008; 317: 549-559
Pearce, LR, Komander, D & Alessi, DR. The nuts and bolts of AGC protein kinases. Nat.
Rev. Mol. Cell Biol 2010; 11: 9-22
Pierce-Shimomura J, Faumont S, Gaston M, Pearson B & Lockery S. The homeobox
gene lim-6 is required for distinct chemosensory representations in C. elegans. Nature
2001; 410:694-698
83
Piggott BJ, Liu J, Feng Z, et al. The neural circuits and synaptic mechanisms underlying
motor initiation in C. elegans. Cell 2011; 147: 922-33
Pirri JK, McPherson AD, Donnelly JL, Francis MM & Alkema MJ. A tyramine-gated
chloride channel coordinates distinct motor programs of a Caenorhabditis elegans escape
response. Neuron 2009; 62(4): 526-38
Pokala N, Liu Q, Gordus A & Bargmann CI. Inducible and titratable silencing of
Caenorhabditis elegans neurons in vivo with histamine-gated chloride channels. Proc
Natl Acad Sci U S A 2014; 111: 2770-5
Purcell J, Robertson AP, Thompson DP & Martin RJ. PF4, a FMRFamide-related
peptide, gates low-conductance Cl- channels in Ascaris suum. Eur J Pharmacol 2002;
456(1-3): 11-7
Ramot D, Johnson BE, Berry TL Jr, et al. The Parallel Worm Tracker: a platform for
measuring average speed and drug-induced paralysis in nematodes. PloS One 2008; 3:
e2208
Ranganathan R, Cannon SC & Horvitz HR. MOD-1 is a serotonin-gated chloride channel
that modulates locomotory behaviour in C. elegans. Nature 2000; 408: 470-475
84
Rao VT, Accardi MV, Siddiqui SZ, et al. Characterization of a novel tyramine-gated
chloride channel from Haemonchus contortus. Mol Biochem Parasitol 2010; 173: 64-68
Rao VT, Forrester SG, Keller K & Prichard RK. Localization of serotonin and dopamine
in Haemonchus contortus. Int J Parasitol 2011; 41: 249-54
Razzaque Z, Heald MA, Pickard JD, et al. Vasoconstriction in human isolated middle
meningeal arteries: determining the contribution of 5-HT1B- and 5-HT1F-receptor
activation. Br J Clin Pharmacol 1999; 47(1): 75-82
Reinitz CA & Stretton AO. Behavioral and cellular effects of serotonin on locomotion
and male mating posture in Ascaris suum (Nematoda). J Comp Physiol 1996; A.178: 655667
Reinitz CA, Herfel HG, Messinger LA & Stretton AO. Changes in locomotory behavior
and cAMP produced in Ascaris suum by neuropeptides from Ascaris suum or
Caenorhabditis elegans. Mol Biochem Parasitol 2000; 111: 185-97
Reinitz CA, Pleva AE & Stretton AO. Changes in cyclic nucleotides, locomotory
behavior, and body length produced by novel endogenous neuropeptides in the parasitic
nematode Ascaris suum. Mol Biochem Parasitol 2011; 180(1): 27-34
85
Rex E, Molitor SC, Hapiak V, et al. Tyramine receptor (SER-2) isoforms are involved in
the regulation of pharyngeal pumping and foraging behavior in Caenorhabditis elegans. J
Neurochem 2004; 91: 1104-15
Reynoldson JA, Behnke JM, Pallant LJ, Macnish MG, Gilbert F, Giles S, Spargo RJ &
Thompson RC. Failure of pyrantel in treatment of human hookworm infections
(Ancylostoma duodenale) in the Kimberley region of north west Australia. Acta Trop
1997; 68: 301-312
Riddle DL, Blumenthal T, Meyer BJ, et al. editors. Section II, cuticle. In: C. elegans II.
2nd edition 1997a
Available: http://www.ncbi.nlm.nih.gov/books/NBK20029/
Accessed: 7 January 2015
Riddle DL, Blumenthal T, Meyer BJ, et al. editors. Section IV, the nematode surface. In:
C. elegans II. 2nd edition 1997b
Available: http://www.ncbi.nlm.nih.gov/books/NBK20022/
Accessed: 7 January 2015
Ringstad N, Abe N & Horvitz HR. Ligand-gated chloride channels are receptors for
biogenic amines in C. elegans. Science 2009; 325(5936): 96-100
86
Rosoff ML, Burglin TR & Li C. Alternatively-spliced transcripts of the flp-1 gene encode
distinct FMRFamide-like peptides in Caenorhabditis elegans. Neurosci 1992; 12: 23562361
Rosoff ML, Doble KE, Price DA & Li C. The flp-1 propeptide is processed into multiple,
highly similar FMRFamide-like peptides in Caenorhabditis elegans. Peptides 1993; 14:
331-338
Ruiz-Lancheros E, Viau C, Walther TN, et al. Activity of novel nicotinic anthelmintics in
cut preparations of Caenorhabditis elegans. Int J Parasitol 2011; 41: 455-61
Schwarz EM, Korhonen PK, Campbell BE, Young ND, Jex AR, Jabbar A, Hall RS,
Mondal A, Howe AC, Pell J, Hoffman A et al. The genome and developmental
transcriptome of the strongylid nematode Haemonchus contortus. Genome Biol 2013;
14(8): R89
Schultz RD, Bennett EE, Ellis EA & Gumienny TL (2014) Regulation of Extracellular
Matrix Organization by BMP Signaling in Caenorhabditis elegans. PLoS One 2014; 9:
e101929
Sheriff JC, Kotze AC, Sangster NC & Martin RJ. Effects of macrocyclic lactone
anthelmintics on feeding and pharyngeal pumping in Trichostrongylus colubriformis in
vitro. Parasitology 2002; 125: 477-484
87
Smith MW, Borts TL, Emkey R et al. Characterization of a novel G-protein coupled
receptor from the parasitic nematode H. contortus with high affinity for serotonin. J
Neurochem 2003; 86: 255-66
Smout MJ, Kotze AC, McCarthy JS & Loukas A. A novel high throughput assay for
anthelmintic drug screening and resistance diagnosis by real-time monitoring of parasite
motility. PloS Negl Trop Dis 2010; 4: e885
Stiernagle T. Maintenance of C. elegans. In: WormBook: The online review of C. elegans
biology 2006
Available: http://www.ncbi.nlm.nih.gov/books/NBK19649/
Accessed: 8 January 2015
Stretton AO. Anatomy and development of the somatic musculature of the nematode
Ascaris. J Exp Biol. 1976; 64(3): 773-88
Summers PJ, Layne RM, Ortega AC, Harris GP, et al. Multiple Sensory Inputs Are
Extensively Integrated to Modulate Nociception in C. elegans. J Neurosci 2015; 35(28):
10331-42
88
Suzuki H, Thiele T, Faumont S, Ezcurra M & Lockery S et al. Functional asymmetry in
Caenorhabditis elegans taste neurons and its computational role in chemotaxis. Nature
2008; 454: 114-117
Thompson DP, Davis JP, Larsen MJ, Coscarelli EM, Zinser EW, Bowman JW,
Alexander-Bowman SJ, Marks NJ & Geary TG. Int J Parasitol 2003; 33(2): 199-208
Thompson AJ, Lester HA & Lummis SC. The structural basis of function in Cys-loop
receptors. Q Rev Biophys 2010; 43(4): 449-99
Waller PJ. International approaches to the concept of integrated control of nematode
parasites of livestock. Int J Parasitol 1999; 29(1): 155-64
Wang SJ & Wang ZW. Track-a-worm, an open-source system for quantitative
assessment of C. elegans locomotory and bending behavior. PloS One 2013; 8: e69653
Wani I, Rather M, Naikoo G, Amin A, Mushtaq S & Nazir M. Intestinal Ascariasis in
Children. World J Surg 2010; 34: 963-8
Welz C, Kruger N, Schniederjans M, et al. SLO-1-channels of parasitic nematodes
reconstitute locomotor behaviour and emodepside sensitivity in Caenorhabditis elegans
slo-1 loss of function mutants. PLoS Pathog 2011; 7: e1001330
89
White WH, Gutierrez JA, Naylor SA, et al. In vitro and in vivo characterization of pamino-phenethyl-m-trifluoromethylphenyl piperazine (PAPP), a novel serotonergic
agonist with anthelmintic activity against Haemonchus contortus, Teladorsagia
circumcincta and Trichostrongylus colubriformis. Vet Parasitol 2007; 146: 58-65
Williamson SM, Robertson AP, Brown L, Williams T, Woods DJ, Martin RJ, Sattelle DB
& Wolstenholme AJ. The nicotinic acetylcholine receptors of the parasitic nematode
Ascaris suum: formation of two distinct drug targets by varying the relative expression
levels of two subunits. PLoS Pathog 2009; 5(7): e1000517
Wolstenholme AJ, Fairweather I, Prichard R, Von Samson-Himmelstjerna G & Sangster
NC. Drug resistance in veterinary helminths. Trends Parasitol 2004; 20: 426-76
Yates DM & Wolstenholme AJ. Dirofilaria immitis: identification of a novel ligandgated ion channel-related polypeptide. Exp Parasitol 2004; 108(3-4): 182-5
Zhang Y, Nash L & Fisher AL. A simplified, robust, and streamlined procedure for the
production of C. elegans transgenes via recombineering. BMC Dev Biol 2008; 8: 119
Zhang Y, MacArthur C, Mubila L & Baker S. Control of neglected tropical diseases
needs a long-term commitment. BMC Med 2010; 8: 67
90