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date: 29 February 2020

Flatworm Neurobiology in the Postgenomic Era

Abstract and Keywords

Flatworm nervous systems comprise central and peripheral components that facilitate coordinated and complex behaviors that are modulated by physiological status and sensory input. The absence of a body cavity in flatworms enhances their dependence on neuronal signaling for intercellular communication. Significant advances have been made in our understanding of the neurobiology of flatworms, largely through the growth in genomic/transcriptomic resources and some progress in the development of functional genomics tools. This chapter describes the “state of the art” of flatworm neurobiology with a primary focus on the recent advances made in parasitic flatworms where progress has been driven by the search for new targets for chemotherapeutics.

Keywords: classical neurotransmitters, neuropeptide, ion channels, G protein–coupled receptors, RNA interference

For several decades, helminth neurotransmission has been recognized as an appealing source of targets for new anthelmintic drugs (Geary et al., 1999; Maule et al., 2002; Mousley et al., 2004; Ribeiro et al., 2005). This appeal stems from two factors: first, the demonstrable importance of the neuromuscular system (NMS) in essential functions such as motility, chemosensation, reproduction, attachment, and feeding; second, many of the core receptors, enzymes, and transporters involved in neurotransmitter biosynthesis and signal transduction are demonstrably “druggable.” That is, they represent protein families against which active compounds already exist in human and/or veterinary medicine. G protein–coupled receptors (GPCRs), ligand-gated ion channels (LGICs), and neurotransmitter transporters are particularly good examples of such targets.

This chapter aims to describe the current state of knowledge regarding the neurotransmitters, receptors, and signaling proteins employed by the flatworm nervous system, in the context of how their structural and functional data can be translated to novel control options for parasites.

In covering the entirety of flatworm neurotransmission, this is necessarily a broad review. Readers interested in more detail on specific areas are directed to reviews on classical transmitters (Ribeiro et al., 2005; Ribeiro, 2015), neurotransmitter transporters (Ribeiro & Patocka, 2013a), neuropeptides (McVeigh et al., 2005b; Marks & Maule, 2010), and anthelmintic drug discovery (Geary, 2010; Geary et al., 2015).

Structure and Function of the Helminth Neuromuscular System

All four classes of phylum Platyhelminthes (Cestoda, Monogenea, Planaria, and Trematoda) display a common neuronal architecture, comprising central and peripheral nervous systems (the definitive description of flatworm neuroanatomy can be found in Halton & Gustafsson, 1996). Typically, the central nervous system (CNS) consists of a relatively primitive anterior brain, composed of paired (p. 220) anterio-lateral ganglia, from which neurons project posteriorly along the paired main nerve cords, laterally across the cerebral commissure, and anteriorly toward the mouth/oral sucker/anterior sense organs. Posterior to the brain, the longitudinal nerve cords are intermittently connected by transverse ring commissures to generate a distinctive orthogonal (ladder-like) appearance (Halton & Gustafsson, 1996). Peripheral motor and sensory neurons (peripheral nervous system, PNS) branch from the CNS, forming fine nerve plexuses that innervate the somatic muscle and surface sensory organs. The nerve cells themselves encompass unipolar, bipolar, and multipolar cells; are typically unmyelinated; and are often found in close association with what are interpreted to be glial cells (Halton & Gustafsson, 1996). Flatworm neurons interact with other cells at synapses that exhibit broad structural diversity. Typical synaptocrine chemical synapses include two-way (nonpolarized) junctions, as well as polarized synapses via which neurons may interact with single or multiple postsynaptic cells. In the case of neuromuscular junctions, nerves may make a traditional synapse onto muscle, or the muscle can reach out to the axon via a cytoplasmic extension. Nonsynaptic release sites exist, where neurosecretory vesicles may be exocytosed from the neuronal membrane directly onto an abutting nerve fiber, fulfiling a neuromodulatory role rather than that of a direct synaptic transmitter.

Flatworms employ classical/small molecule and peptidergic neurotransmitters, both of which are widely distributed throughout the CNS and PNS (Ribeiro et al., 2005; McVeigh et al., 2005b). These two neurotransmitter classes are generated by distinct biosynthetic means, but their synaptic release, receptor interactions, and signaling pathways employ relatively similar mechanisms. Classical transmitters are produced by biosynthetic enzymes present in the cytosol of a presynaptic nerve terminal, with each transmitter synthesized by a discrete set of enzymes (Fig. 8.1), before being loaded into secretory vesicles by transmitter-specific transporter pumps. Neuropeptides are produced via the protein synthesis pathway, as single or multiple copies of related peptides within larger precursor proteins (prepropeptides). Mature peptides are subsequently excised from the prepropeptide, ahead of their posttranslational processing. This processing occurs entirely within the secretory pathway of the endoplasmic reticulum (ER) and Golgi, resulting in vesicles filled with mature, processed peptides. Secretory vesicles of either classical or peptidergic origin accumulate at the presynaptic membrane, ready for release upon receipt of an appropriate signal. Generally, the arrival of an electrical impulse triggers the influx of calcium (Ca2+) into the cytosol through voltage-operated Ca2+ channels (VOCCs). Subsequently increased cytosolic [Ca2+] causes conformational changes in SNARE-like docking proteins (Hong & Lev, 2014), resulting in membrane fusion and vesicular exocytosis. Transmitters released into the synapse interact with receptors present on the postsynaptic membrane. Flatworm neurotransmitters most typically employ one of two types of receptors—ligand-gated ion channels (LGICs) and G protein–coupled receptors (GPCRs).

Among classical neurotransmitters there are examples of both LGIC- and GPCR- mediated signal transduction, while all the flatworm neuropeptides described to date appear to signal through GPCRs. Regardless of receptor type, ligands must be removed from the synapse following receptor activation before further signals can be transmitted, in a process known as signal termination. Peptidergic- and cholinergic-mediated signals are terminated by enzymes that destroy the transmitter within the synapse. All remaining classical transmitters are removed from the synapse by specific transporter pumps, which actively transport transmitters back into neurons or glial cells for metabolic breakdown or reuse.

Classical Neurotransmitters

Experiments performed from the 1960s onward demonstrated the presence in flatworms of a complement of now well-established classical/small-molecule neurotransmitters, including acetylcholine, dopamine, γ-aminobutyric acid, glutamate, histamine, noradrenaline, octopamine, and serotonin, all of which have been directly or indirectly detected in several flatworm species and/or have been localized to neuronal tissue (Ribeiro et al., 2005). Many also have demonstrable activity when added to whole flatworms or neuromuscular preparations in vitro. Although no existing drugs target classical neurotransmission in flatworms, the potential of this system as a source of therapeutic targets is highlighted by the many antinematode anthelmintics that target classical transmitter receptors. Dichlorvos, levamisole, morantel, piperazine, pyrantel, macrocyclic lactones, paraherquamide, and the amino acetonitrile derivatives (AADs) all act upon receptors or enzymes associated with classical neurotransmission (McVeigh et al., 2012). (p. 221)

Flatworm Neurobiology in the Postgenomic Era

Figure 8.1 Biosynthetic pathways of classical neurotransmitters. Precursors and enzymes responsible for biosynthesis of the classical transmitters present in flatworms are shown. Accession numbers are presented for enzymes that have been cloned or reported in genome publications. RNA interference (RNAi) phenotypes are also summarized where available. References: DjChAT (Nishimura et al., 2010); Tyrosine hydroxylase (Hamdan & Ribeiro, 1998; Hu et al., 2011); DjGAD (Nishimura et al., 2008a); DjTBH (Nishimura et al., 2008b); and SmTPH (Hamdan & Ribeiro, 1999).

(p. 222) Acetylcholine

Acetylcholine (ACh) is the primary excitatory transmitter at neuromuscular junctions of vertebrates and most invertebrates. It has species-specific excitatory or inhibitory neuromuscular impacts in flatworms, triggering muscle contraction in planarian preparations (Moneypenny et al., 2001; Blair & Anderson, 2009), while inhibiting myoactivity in trematode and cestode assays (Pax et al., 1984; Sukhdeo et al., 1984, 1986; Thompson et al., 1986; McKay et al., 1989; Day et al., 1996; Maule et al., 2009). In addition to its core neuromuscular role, there is also evidence that ACh is involved in nutrient acquisition at the host-parasite interface, since nicotinic ACh receptors (nAChRs) and acetylcholinesterase (AChE) are concentrated on the tegumental surface of adult male schistosomes. These cholinergic signaling elements are linked to schistosome glucose transporters, enabling glucose uptake from host blood, triggered by typical circulating concentrations of ACh. Glucose transport is ablated by both nAChR antagonists and AChE inhibitors (Camacho & Agnew 1995). This highlights an additional opportunity for anticholinergic compounds to target schistosomes; by targeting these surface-exposed elements, it may be possible to use preexisting pharmacology to interfere with parasite energy metabolism or glycolysis.

Biosynthesis of ACh from choline and acetyl coenzyme A is catalyzed by choline acetyltransferase (ChAT) (Fig. 8.1), with signaling terminated following receptor activation by the hydrolytic action of AChE. AChE activity can be detected histochemically, providing a convenient marker for cholinergic synapses—indeed, cholinesterase histochemistry provided the first evidence of ACh in flatworms (Graft & Read, 1967; Halton, 1967). ChAT, ACh’s single biosynthetic enzyme, has been poorly studied in flatworms, with only a single publication reporting cloning and functional analysis of this protein from the planarian Dugesia japonica (Nishimura et al., 2010). Nishimura et al. (2010) employed RNA interference (RNAi) to silence the DjChAT gene, yielding an inhibited motility phenotype that phenocopied the impacts of nicotinic ACh receptor antagonist drugs. This effect is consistent with DjChAT’s involvement in the synthesis of the excitatory ACh.

ACh is a ligand at both nicotinic (LGIC) and muscarinic (GPCR) receptors. Muscarinic ACh receptor (mAChR)-like sequences have been identified in cestode, planarian, and trematode genomes (Zamanian et al., 2011; Tsai et al., 2013; Campos et al., 2014). One mAChR (also termed GAR: G protein–coupled acetylcholine receptor) has been functionally expressed and characterized from S. mansoni (MacDonald et al., 2015) (Fig. 8.2). SmGAR expressed in yeast showed basal constitutive activity, with further activation triggered by the addition of ACh, but not tyramine, histamine, or glutamate, suggesting that SmGAR is a specific ACh receptor. RNAi analysis indicated that SmGAR has a role in neuromuscular coordination, since silencing the receptor inhibited the motility of larval S. mansoni (MacDonald et al., 2015) (Fig. 8.2).

Vertebrate nAChRs are invariably selective for cations, triggering membrane depolarization and cellular excitation when activated. The classic example of ACh-nAChR signaling is in the contraction of skeletal muscle at the vertebrate neuromuscular junction. Three nicotinic ACh receptor (nAChR) subunits have been cloned and localized in S. haematobium, one of which (ShAR2beta) was considered divergent based on the amino acids lining the ion pore (Bentley et al., 2004, 2007; MacDonald et al., 2014). ShAR1alpha and ShAR2beta are expressed in the S. haematobium tegument and musculature, respectively, but neither subunit formed functional channels, either alone, in combination, or coupled with a vertebrate alpha7 subunit, in Xenopus oocytes (Bentley et al., 2004). Given the availability of extensive nicotinic pharmacology that could potentially delineate parasite nAChRs from those of the host, the inability to functionally express these schistosome channels could be considered a missed opportunity for drug discovery. However, the presence of at least 12 nicotinic subunits in the S. mansoni genome (including homologues of the three S. haematobium subunits described earlier) (Berriman et al., 2009) could stimulate further efforts in this regard.

Invertebrates exhibit some interesting diversification of the ACh-gated ion channel complement; in addition to cation-selective nAChRs, they also possess anion (Cl)-selective ACh-gated channels (ACCs). By transmitting negative charge, ACCs trigger cellular hyperpolarization and are therefore associated with inhibitory signaling. In molluscs, and nematodes (Putrenko et al., 2004; van Nierop, 2005; Beech et al., 2013) ACC subunits are defined by the presence of a Pro-Ala anion-selective motif in the M2 domain that lines the channel pore. MacDonald et al. (2014) demonstrated the presence of this motif in five S. mansoni nAChR subunits. (p. 223) (p. 224) The anion-selective motif is also found in ShAR2 beta, the previously identified divergent nAChR subunit from S. haematobium (Bentley et al., 2007). Silencing ACC subunits in schistosomula yielded a hyperactive phenotype, consistent with the abolition of inhibitory signaling (Fig. 8.2). Heterologous expression of one subunit, SmACC-1, yielded a functional homomeric channel in HEK-293 cells that mediated anion influx upon exposure to nicotine and other cholinergic agonists. This response was inhibited by the nicotinic antagonist, D-tubocurarine, but not by other classical nicotinic antagonists (mecamylamine and atropine) (MacDonald et al., 2014). The tissue expression patterns of SmACC subunits have not yet been described, making it difficult to ascertain whether the motility impacts of SmACC RNAi are the result of silencing neuron- or muscle-expressed receptors. Nevertheless, these data suggest that ACC channel subunits may be responsible for at least some of the myoinhibitory effects of ACh on schistosomes (Day et al., 1996). Comparative studies on the presence and/or relative distribution and function of ACC in the planarian NS would be informative in the context of ACh’s myoexcitatory actions in these turbellarians.

Flatworm Neurobiology in the Postgenomic Era

Figure 8.2 Deorphanized flatworm receptors. Fourteen flatworm-derived receptors have been deorphanized in heterologous expression systems or by RNA interference (RNAi) (GtSER1 and GtNPR1 only). All are activated by classical transmitters except GtNPR1, which is a neuropeptide receptor. The ATP-gated channels (SchP2X and SmP2X), an acetylcholine-gated chloride channel (SmACC), and the glutamate gated chloride channels (SmGluCl) are ligand-gated ion channels; all of the remaining receptors are seven-transmembrane G protein–coupled receptors (GPCRs). Receptors are colored according to activity, where red represents a receptor with inhibitory function, green indicates excitatory function, and gray indicates unknown function. Heterologous expression systems, signaling pathways, function, and RNAi phenotypes are indicated where known. References: SchP2X (Agboh et al., 2004); SmP2X (Raouf et al., 2005); SmACC1 (MacDonald et al., 2014); SmGAR (MacDonald et al., 2015); SmD2 (Taman & Ribeiro, 2009); SmGPR3 (El-Shehabi et al., 2012); SmGluCl-1,-2,-3 (Dufour et al., 2013); SmGluR (El-Shehabi et al., 2009); SmGPCR (Fadi F Hamdan et al., 2002); SmGPR2 (El-Shehabi & Ribeiro, 2010); Sm5HTR (Patocka et al., 2014); Dj5HT7 (Nishimura et al., 2009); GtSER1 (Zamanian et al., 2012); and GtNPR1 (Omar et al., 2007; Zamanian et al., 2012).

ACh signaling is terminated by the hydrolytic action of AChE present within cholinergic synapses. The potential of AChE as a drug target is highlighted by the success of organophosphate pesticides, which function as AChE inhibitors. Although now banned due to toxicity concerns, these compounds were extremely effective weapons against arthropods, nematodes, and other agricultural pest species, in which they caused muscular paralysis. AChEs from three Schistosoma spp. have been functionally expressed (Bentley et al., 2003, 2005). Importantly, these show structural differences from mammalian enzymes that might enable the development of compounds with selectivity between helminth and host AChE variants.

Catecholamines—Dopamine, Norepinephrine, and Epinephrine

The catecholamine biosynthetic pathway is responsible for the production of dopamine, norepinephrine, and epinephrine. Catecholamines mediate a vast amount of CNS signaling in mammals, with roles in motor function, learning, and memory (Kobayashi, 2001). Dopamine has similarly central roles in invertebrates; in Drosophila, dopamine controls basal locomotion, as well as more complex functions in circadian rhythm, learning, and courtship behavior (Yamamoto & Seto, 2014). Dopamine is also involved in the regulation of molluscan feeding behavior (Baxter & Byrne, 2006). There is limited evidence for the presence of norepinephrine and epinephrine in invertebrate phyla, with the phenolamines (tyramine and octopamine) considered their invertebrate equivalents (Xu et al., 2015). Nevertheless, there is some evidence for the presence of both dopamine and noradrenaline in flatworms. Both transmitters, and their biosynthetic activities, have been identified in cestodes, planaria, and trematodes (Ribeiro et al., 2005). Dopamine and norepinephrine have distinct neuromuscular effects when applied to different parasitic flatworm species, including circular muscle relaxation (S. mansoni; Hillman & Senft, 1973; Tomosky et al., 1974; Mellin et al., 1983; Pax et al., 1984), muscle contraction (D. merlangi; A. G. Maule et al., 2009), and dual excitatory/inhibitory effects (F. hepatica; Holmes & Fairweather, 1984; Davis & Stretton, 1995; Ribeiro et al., 2005).

Dopamine is synthesized in a two-step reaction from tyrosine (Fig. 8.1), which is first converted to L-Dopa by tyrosine hydroxylase; L-dopa is subsequently converted to dopamine by dopa decarboxylase. Full-length, enzymatically functional examples of tyrosine hydroxylase have been cloned from schistosomes (Hamdan & Ribeiro 1998; Hu et al., 2011). Dopa decarboxylase (also known as aromatic L-amino acid decarboxylase) has not yet been reported in any flatworm, but basic searches of available flatworm genomes support this enzyme’s existence in S. mansoni (Fig. 8.1). Norepinephrine is synthesized from dopamine by dopamine β hydroxylase (DBH). DBH has currently only been described in Schmidtea mediterranea, where it was used as a marker for transcription factors necessary for regeneration of noradrenergic neurons (Scimone et al., 2014). Figure 8.1 shows that a DBH homologue also appears in the S. mansoni genome. In mammals, epinephrine is produced from norepinephrine by phenylethanolamine N-methyltransferase (PNMT; Fig. 8.1). Neither epinephrine, nor PNMT activity, has been reported in any flatworm.

Dopamine signaling is transduced through GPCRs. In mammals, these receptors are classified into two families (D1 and D2 receptors), which themselves encompass five subtypes (the D1 family contains D1 and D5 subtypes; the D2 family comprises D2, D3, and D4 subtypes). D1 and D2 families are distinguished by signal transduction mechanism, where D1 receptors stimulate cAMP (p. 225) production through G protein Gαs/olf subunits, while D2 receptors inhibit cAMP through Gαi/o (Beaulieu & Gainetdinov, 2011). In flatworms, two dopamine-activated GPCRs have been identified from S. mansoni (SmD2 and SmGPR3; Fig. 8.2). SmD2 clusters phylogenetically with D2 sequences, suggesting D2-like sequence identity, but lacks the typical pharmacology of mammalian D2 receptors when challenged with dopaminergic agonists and antagonists. Most notable is the effect of apomorphine, a mammalian dopaminergic antagonist which acts as an agonist at heterologously expressed SmD2 (Taman & Ribeiro, 2009). SmGPR3 is one representative of a schistosome-specific clade of biogenic amine GPCRs lacking homologues in other genera (El-Shehabi et al., 2012), and it was detected throughout the schistosome nervous system in a pattern similar to that of dopamine (El-Shehabi et al., 2012). When exposed to catecholamines, heterologously expressed SmGPR3 was most potently activated by dopamine. When challenged with classical biogenic amine antagonists, SmGPR3’s profile did not resemble that of any known mammalian dopaminergic receptor. Reminiscent of apomorphine’s impact on SmD2, spiperone, a mammalian D2 antagonist displayed agonism of SmGPR3.

Catecholamines are removed from the synapse by transporter pumps. A single dopamine transporter (DAT) has been cloned and functionally characterized from S. mansoni. Recombinant SmDAT showed dopamine transport activity, but interestingly, displayed pharmacology more akin to the human norepinephrine transporter because of its low affinity for known DAT inhibitors, including cocaine (Larsen et al., 2011). This pharmacological difference may permit selective targeting of this transporter for therapy.

γ-Aminobutyric Acid

γ-aminobutyric acid (GABA) is synthesized from glutamate by glutamate decarboxylase (GAD; Fig. 8.1). GABA itself has been identified in high performance liquid chromatography (HPLC) analyses from cestode, planarian, and trematode nervous systems (Ribeiro et al., 2005). GAD biosynthetic activity has been described in adult S. mansoni (Mendonça-Silva et al., 2004). GAD has also been described in the planarian, D. japonica (DjGAD) (Nishimura et al., 2008a), as a marker for GABAergic neurons, where expression of both DjGAD mRNA and protein were localized to neurons of the brain and pharynx. Nishimura et al. (2008a) also used RNAi to silence DjGAD, triggering a reduction in detectable GABA, alongside an aberrant photosensitivity phenotype. DjGAD-knockdown ablated the negative phototaxis seen in wild-type worms, although their locomotion remained normal. These data suggest that GABAergic transmission is associated with planarian photosensitivity, but not motility.

GABA operates through two types of receptor—GABAA (GABA-gated Cl channels (Vogt, 2015) and GABAB (GABA-activated GPCRs; Brown et al., 2015). Neither of these receptor types has been cloned or characterized from any flatworm. Indeed, Dufour et al. (2013) stated that they found no evidence for schistosome GABA receptors during database searches that led to their discovery of the first S. mansoni glutamate-gated Cl channels. Similarly, three GPCR discovery papers make no mention of GABAB receptors in the S. haematobium, S. mansoni, and S. mediterrannea genomes (Zamanian et al., 2011; Tsai et al., 2013; Campos et al., 2014). There is some pharmacological evidence for GABAA receptors in schistosomes. Modulation of schistosome motility by picrotoxin, a GABAA receptor antagonist, implicates ionotropic GABA receptors in the control of schistosome motility (D. L. Mendonça-Silva et al., 2004), and there is evidence that benzodiazepine, a positive allosteric modulator of GABAA receptors, binds to schistosome extracts (Noël et al., 2007). These data suggest that GABAA receptors may be present in schistosomes, although their presence remains to be confirmed by cloning and functional characterization. No synaptic GABA transporters (GAT) have been formally identified in flatworms, but there is phylogenetic evidence for the presence of three GAT-like sequences in S. mansoni (Ribeiro & Patocka, 2013a).


Glutamate is synthesized from glutamine, by glutaminase (Fig. 8.1). This amino acid is ubiquitous throughout nature due to its obvious importance for protein synthesis, but glutamate is also an important neurotransmitter in both vertebrates and invertebrates. In vertebrates, glutamate is an excitatory transmitter that signals through both ionotropic glutamate receptors (iGluRs—LGICs) and metabotropic glutamate receptors (mGluRs—GPCRs). Invertebrates display additional functionality where glutamate may also trigger inhibitory signaling through glutamate-gated Cl (GluCl) channels (Wolstenholme, 2012).

(p. 226) Ionotropic Glu receptors are composed of five subunit types: (i) AMPA (α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid); (ii) NMDA (N-methyl-D-aspartate); (iii) Kainate; and (iv) Delta subunits δ1 and δ2, termed “orphan subunits” because they have not yet been shown capable of forming a functional channel (Brockie, 2006). GluCl, AMPA, NMDA, Kainate, and Delta receptors exist in both vertebrates and invertebrates, (Brockie, 2006; Collingridge et al., 2009), while the GluCls are restricted to protostome invertebrates (Wolstenholme, 2012). Glutamate signaling is additionally transduced through mGluRs, of which there are eight subtypes (mGluR1-8) in mammalian systems (Niswender & Conn, 2010), and only one in Drosophila (Schoenfeld et al., 2013).

While none of the excitatory iGluR subunits have been cloned from flatworms, there is some pharmacological evidence for their presence. Kainate receptor pharmacology has been reported in schistosomes, radioactively labeled kainate binds to subcellular fractions of adult male S. mansoni, while addition of kainate to whole schistosomes induces a “corkscrew” motility phenotype (Mendonça-Silva et al., 2002). Similarly, both NMDA and AMPA induce seizure-like modulations of motility in planaria, similar to their impacts in mammalian models (Rawls et al., 2009). Four GluCl subunits have been cloned from S. mansoni (Dufour et al., 2013), three of which produced glutamate-gated Cl-permeable channels when expressed in Xenopus oocytes (Fig. 8.2).

Several mGluRs are present in flatworm genomes—nine have been identified in S. mediterranea, three in Echinococcus multilocularis, and two each in S. mansoni and S. haematobium (Zamanian et al., 2011; Tsai et al., 2013; Campos et al., 2014). One of these S. mansoni receptors was cloned, functionally expressed, and localized to the central and peripheral nervous systems of adult and larval worms, and the adult female reproductive tract (Taman & Ribeiro, 2011b). This receptor was activated by glutamate, and it was unresponsive to GABA (Fig. 8.2). Importantly, the receptor’s response profile to classical glutamatergic modulators suggests that it has quite different pharmacology from mammalian mGluRs, indicating its potential as a target for schistosome-specific drugs. An additional truncated mGluR has been reported in S. mansoni (SmGBP) (Taman & Ribeiro, 2011a), consisting of the extracellular N-terminal ligand binding domain, but lacking the heptahelical transmembrane domain that is characteristic of these receptors. SmGBP binds glutamate in radiolabeled glutamate competition assays. Expressed solely in the surface tubercles of adult male worms, Taman and Ribeiro (2011b) hypothesize a role in host-parasite signaling for SmGBP, although since it lacks the structural features required for G protein interaction, this truncated GluR must operate via another, potentially novel, signaling mechanism. Truncated GluRs were identified in other flatworms but have not been described in other metazoans (Taman & Ribeiro, 2011a). Metabotropic glutamate receptors are known to form functional dimers, where such interactions are essential for signaling (Pin & Acher, 2002; Kniazeff et al., 2004; Kammermeier & Yun, 2005). Immunoprecipitation experiments showed that SmGBP forms noncovalent dimers at neutral pH; Taman and Ribeiro hypothesized that SmGBP may regulate glutamatergic signaling by interacting/dimerizing with full-length mGluRs.

In cestode preparations, glutamate addition triggers a range of neuromuscular impacts, including contraction of muscle strips, increased electrical signaling in nerve tissue, and production of inositol triphosphate (IP3) second messengers (Ribeiro et al., 2005), the latter of which implicates the involvement of mGluRs in cestode glutamatergic signaling. The impact of glutamate on other flatworms is less compelling (Ribeiro et al., 2005). Aside from the aforementioned reports of kainate activity in schistosomes (Mendonça-Silva et al., 2002) and AMPA/NMDA impacts on planarian motility (Rawls et al., 2009), there is no strong evidence for the direct involvement of glutamate receptors in the motility of monogeneans, planarians, or trematodes. In the latter case, although glutamate does cause isolated schistosome muscle fibers to contract, this is thought to be an electrogenic, rather than receptor-mediated effect (Miller et al., 1996).

Little is known of flatworm synaptic glutamate transporters (GLTs), aside from a bioinformatics analysis that predicts a single GLT in the S. mansoni genome, of the solute carrier (SLC1) family (Ribeiro & Patocka 2013a).


Histamine biosynthesis is performed from histidine by histidine decarboxylase (Fig. 8.1). Although some flatworms are known to possess this enzyme and produce endogenous histamine, others are thought to obtain histamine from their host (El-Shehabi & Ribeiro, 2010). Histamine (p. 227) acts exclusively through GPCRs, which in humans comprise four families, H1–H4 (Stark, 2007). Exogenous applications of histamine have generally positive impacts on motility of live, intact cestodes and trematodes, and some H1 antagonists (H2, H3, and H4 blockers have no effects) have correspondingly inhibitory effects on motility (Ribeiro et al., 2005). Histamine lacks impacts on the motility of flatworm neuromuscular assay preparations (i.e., exposed neuromuscular assays in the form of muscle strips, trimmed worms, or dispersed muscle fibers). In line with these data, it has been suggested that rather than acting as an endogenous signaling molecule, histamine’s stimulation of parasite motility may in fact enable the parasite to respond to the presence of host histamine, levels of which increase following infection with schistosomes (Ribeiro et al., 2005). Given histamine’s role in host immune signaling, such ability would be advantageous, particularly for blood-dwelling schistosomes. Consistent with this hypothesis, one of the two known schistosome histamine receptors (SmGPCR, Fig. 8.2; Hamdan et al., 2002) localizes to the tegument of schistosomula, adult male and intramolluscan sporocyst forms of S. mansoni. The second S. mansoni histamine receptor (SmGPR2, Fig. 8.2; El-Shehabi & Ribeiro, 2010) localizes to the nervous system, suggesting an endogenous neuromuscular role. Both of these GPCRs are activated by histamine in heterologous expression systems (Fig. 8.2). Profiling SmGPR2 with classical histamine antagonists (El-Shehabi & Ribeiro, 2010) reveals distinct pharmacology from mammalian histamine receptors, hinting at the ability to specifically target this receptor.

Synaptic inactivation/removal of histamine occurs by an unknown mechanism, since no known biogenic amine transporter has been shown to exhibit histamine transport activity. Histamine reuptake may instead be performed by unselective organic cation transporters of unknown identity (Ogasawara et al., 2006; Ribeiro & Patocka, 2013b).

Nitric Oxide

Nitric oxide (NO), a gaseous signaling molecule in vertebrates (Ignarro, 1990), has been indirectly identified in flatworms based on the presence of its biosynthetic enzyme, nitric oxide synthase (NOS). NOS synthesizes NO from L-arginine (Fig. 8.1). NO signals via activation of soluble guanylate cyclase, which catalyzes the production of a cyclic GMP (cGMP) second messenger that primarily activates protein kinase G. In vertebrates, this signaling axis performs neurotransmission, as well as regulating aspects of cardiovascular and renal function (Bian & Murad, 2014). NOS, as visualized by histochemical staining of nicotinamide adenine dinucleotide phosphate (NADPH) diaphorase activity (Hope et al., 1991), localizes to neurons in both cestodes and trematodes (Ribeiro et al., 2005). Localization of NOS in adult schistosomes additionally identified NOS immunoreactivity in the surface-exposed gut and tegument (Kohn et al., 2001). Expression of NOS at the parasite surface may suggest a role for NO signaling at the host-parasite interface (Da’dara & Skelly, 2011). Host-derived, NO-mediated signals may be received by schistosomes, given that exogeous NO can affect schistosome transcriptional profiles (Messerli et al., 2006). At present there are no data regarding NO’s function in the flatworm NS or in the parasite’s ability to influence the host environment.

Phenolamines—Tyramine and Octopamine

Phenolaminergic transmitters comprise tyramine and octopamine. Both of these transmitters are synthesized from a single sequential pathway (Fig. 8.1) in which tyramine is produced from tyrosine by tyrosine decarboxylase, which itself is subsequently converted to octopamine by tyramine ß-hydroxylase (TBH). Tyramine and octopamine are considered the invertebrate equivalent of the vertebrate adrenergic transmitters (epinephrine and norepinephrine) (Xu et al., 2015).

Tyramine is an established neurotransmitter in insects (Roeder et al., 2003; Roeder, 2005). Its existence in flatworms is implied by the presence of tyrosine decarboxylase activity in cestodes (Ribeiro & Webb, 1983) (Fig. 8.1). However, since tyramine is also an essential intermediate in octopamine biosynthesis, and its neuro-/myoactivity in any flatworm assay has yet to be demonstrated, its status as an endogenous flatworm neurotransmitter remains uncertain. The occurrence of octopamine in flatworms is similarly implied by the presence of TBH activity in Hymenolepis diminuta extracts (Ribeiro & Webb, 1983). TBH has also been cloned from the planarian Dugesia, where knockdown reduced the amount of HPLC-detectable octopamine in tissue extracts (Nishimura et al., 2008b). Nishimura et al. are the only authors to have performed RNAi on any aspect of the phenolaminergic signaling pathway, and it is notable that they reported no aberrant behavior or motility phenotypes associated with TBH ablation. This may suggest that octopamine, in planarians at least, is not involved in neuromuscular function, or at most has a subtle role that does (p. 228) not obviously influence gross motility or behavior. Octopamine and tyramine both act via GPCRs in invertebrates (Farooqui, 2007; Lange, 2009), although homologues of these receptors have not yet been reported in flatworms. Neither have any synaptic phenolamine transporters been identified in flatworms, although one of the S. mansoni biogenic amine transporters has been tentatively annotated as an octopamine transporter (Caveney et al., 2006; Ribeiro & Patocka, 2013b).


Serotonin (5-hydroxytryptamine; 5HT) is the best studied (in terms of publication volume) of all flatworm classical neurotransmitters. It has been identified in all four platyhelminth classes and appears abundant throughout the entire nervous system (Ribeiro et al., 2005). Biosynthesis of 5HT involves a two-step reaction from tryptophan (Fig. 8.1), which is first converted to 5-hydroxytryptophan by tryptophan hydroxylase (TPH), from which 5HT is produced by L-aromatic amino acid decarboxylase (L-AADC). There is biochemical evidence for both of these enzymes throughout phylum Platyhelminthes (Ribeiro et al., 2005). An S. mansoni homologue of TPH (SmTPH) has been cloned and functionally characterized in vitro (Hamdan & Ribeiro, 1999). Although the human and schistosome enzymes are biochemically similar, the mammalian enzyme is difficult to study in vitro due to extreme instability, whereas SmTPH is stable at 37°C. Consistent with 5HT’s constitutive expression, SmTPH transcripts are detectable in all life stages (Hamdan & Ribeiro, 1999; Boyle et al., 2003). L-AADC (also known as Dopa decarboxylase, where it is also involved in dopamine synthesis—see earlier) has not yet been reported in any flatworm.

5HT is considered the flatworm primary excitatory neurotransmitter. In in vitro assays, exogenously applied 5HT stimulates motility of whole worms and causes muscle contraction in trimmed worms and muscle strips (Ribeiro et al., 2005). On dispersed muscle fibers, 5HT has distinct impacts between schistosome- and planarian-derived preparations. Muscle fibers from the planarian Procerodes littoralis contract when directly exposed to 5HT, in a concentration-dependent manner, while preincubation in serotonin blunts the contractile responses of other myoactive agents, such as FMRFamide-related neuropeptides (Moneypenny et al., 2001). In contrast, serotonin does not cause S. mansoni fibers to contract, but it is a prerequisite for depolarization-induced contraction of schistosome-derived fibers (Day et al., 1994). Although technical differences between these two assays (that employ distinct methods in liberating cells from each species) cannot be ruled out as an explanation for the apparent disparity, these data could indicate the existence of species-specific differences in flatworm serotoninergic neuromuscular physiology. There is also ample evidence for serotoninergic signaling having an important role in platyhelminth developmental processes. As well as evidence linking 5HT and 5HT receptors with planarian cellular regeneration (see Ribeiro et al., 2005, for review), serotonin similarly stimulates the differentiation of protoscoleces toward metacestodes in Echinococcus spp. cestodes (Camicia et al., 2013). The lack of a body cavity and a circulatory system in acoelomate flatworms designates the nervous system as the primary carrier of developmental cues. As such, neurotransmitters could well have developmental signaling roles in addition to those responsible for controlling neuromuscular and sensory functions. We would anticipate similar roles in the control of regeneration and cellular renewal for other transmitters as well as 5HT, although this hypothesis has not been adequately tested.

Mammalian 5HT receptors comprise seven major types (5HT1–7); one (5HT3) is an LGIC, whereas the remainder are GPCRs (D. E. Nichols & Nichols, 2008). The GPCR subtypes are distinguished by signal transduction pathway, where 5HT1 and 5HT5 receptors signal through Gαi/o, by inhibition of cAMP, 5HT2 stimulates IP3 synthesis through Gαq/11, while 5HT4, 6, and 7 stimulate cAMP production via Gαs. Invertebrate 5HT receptor families appear somewhat less complex; C. elegans, D. melanogaster, and Apis mellifera all possess 5HT1, 2, and 7 orthologues (Dernovici et al., 2007; Blenau & Thamm, 2011). C. elegans appears unique in also employing a 5HT-gated Cl channel known as MOD-1 (Ranganathan et al., 2000). Three 5HT-activated GPCRs have been functionally described in flatworms (Fig. 8.2). The first, from D. japonica, was activated by 5HT when functionally expressed in Xenopus oocytes. Dj5HT7 (named for its homology to mammalian 5HT7), signaled through a Gαs-compatible chimeric G protein and did not respond to other neurotransmitters aside from 5HT (Nishimura et al., 2009). A second planarian (Dugesia tigrina) 5HT receptor was identified using a novel, RNAi-guided approach to deorphanization (Zamanian et al., 2012). By performing RNAi on individual receptors alongside a (p. 229) second messenger assay as a post-RNAi phenotypic readout, Zamanian et al. demonstrated that DtSER-1 knockdown reduced the levels of 5HT-stimulated cAMP generated by tissue homogenates. This novel methodology illustrated that DtSER-1 was activated by serotonin, without the need to clone and functionally express the receptor in a heterologous system. The latter, more traditional method was used by Patocka et al. (2014) to deorphanize a 5HT7-like receptor from S. mansoni (Sm5HTR). Sm5HTR is expressed throughout the CNS and PNS, and it is involved in parasite motility. The receptor specifically appears involved in transducing excitatory neuromuscular signals, since Sm5HTR RNAi drastically reduces motility in both larval and adult schistosomes.

Serotonin signal termination occurs by physical removal of 5HT from the synapse, a process performed by serotonin-specific reuptake transporter (SERT) pumps. These pumps are pharmacologically important targets for commonly prescribed antidepressant drugs, including specific serotonin reuptake inhibitors (SSRIs), such as fluoxetine and citalopram, and tricyclic antidepressants, such as imipramine (Gether et al., 2006). The druggability of these transporters, alongside their involvement in demonstrably important neuromuscular functions, highlights their obvious appeal as anthelmintic targets. One SERT has been identified and functionally characterized in S. mansoni (Patocka & Ribeiro, 2007, 2013; Fontana et al., 2009; Ribeiro & Patocka, 2013b). SmSERT is represented by two isoforms. While sensitive to many of the drugs that act on human SERT, the relative potencies of these drugs are quite different on helminth versus host transporters. This indicates the presence of distinct pharmacology that could be exploited to develop parasite-specific SERT-targeting drugs.


Since the 1950s there have been suggestions that ATP was released by neurons following electrical stimulation (Mutafova-Yambolieva & Durnin, 2014). There is now an appreciation that purinergic signaling is involved in many physiological processes, including neurotransmission. Within the mammalian brain, purinergic signaling is responsible for neuromodulation, synaptic plasticity, and even fast synaptic transmission (Burnstock, 2013), processes that are mediated by two types of purinergic receptor—P2X trimeric ionotropic receptors (Wang & Yu, 2016) and P2Y GPCRs (von Kügelgen & Hoffmann, 2015). Following receptor interaction, ATP hydrolysis and signal termination is performed by one of eleven ecto-nucleotidases (Zimmermann & Braun, 1999; Williams, 2002). Limited research attention has been paid to the purinergic signaling components of flatworms. The only evidence for these pathways comes from two reported P2X receptors in S. mansoni (Agboh et al., 2004; Raouf et al., 2005). Since the expression profiles of these receptors remain unknown, their functions remain similarly cryptic. Aside from neurotransmission, schistosome P2X receptors could conceivably be surface expressed and responsible for detecting host ATP in some aspect of host-parasite communication. Clearly, further research is required to understand the importance, if any, of purinergic neurotransmission in flatworms.


Classical neurotransmitters are clearly involved in important aspects of platyhelminth neuromuscular function, and their receptors and synaptic transporters represent validated anthelmintic targets, as evidenced by the RNAi and pharmacological phenotypes detailed earlier. Arguably as significant in terms of function is the peptidergic arm of the flatworm nervous system. Immunocytochemical studies demonstrate that the extent of neuropeptide immunoreactivity outstrips that of classical transmitters in flatworm neuronal elements (Halton & Maule, 2004), and all neuropeptides tested on flatworm bioassays have invariably been biologically active (exhibiting either myoactivity or stimulation of signaling activity). Allied to clear differences in peptide signaling and function between parasites and host organisms, these factors argue in favor of flatworm peptidergic signaling as a potential source of targets for new anthelmintic drugs (Mousley et al., 2005; McVeigh et al., 2012). Although well characterized in terms of peptide distribution, physiology, and pharmacology, the flatworm neuropeptidergic nervous system has only in recent years benefited from improved sequence datasets, which have allowed the first characterizations of the genes encoding putative neuropeptides, peptide receptors, and associated signal transduction proteins. Here, we will summarize current understanding of the diversity and function of flatworm neuropeptides and their biosynthetic and signaling elements.

Neuropeptide Biosynthesis—A Summary

Helminth neuropeptide synthesis has been extensively reviewed elsewhere, and the reader is referred (p. 230) to these reviews for graphical representations of the helminth neuropeptide biosynthetic pathway (McVeigh et al., 2005a, 2005b; Husson & Schoofs, 2007a, 2007b; Li & Kim, 2008; Geary, 2010).

Neuropeptides are transcribed and translated from their encoding genes on precursor proteins, known as prepropeptides, that may incorporate single or multiple copies of peptides, with the latter often bearing related motif sequences. Prepropeptides display an N-terminal secretory signal peptide sequence that targets the protein into the cellular secretory pathway. During ribosomal translation, this sequence is the first part of the nascent protein to emerge from the ribosome, enabling its binding to a signal recognition particle (SRP). The SRP translocates the growing protein and its ribosome to the ER, where the protein continues to be synthesized through a translocon (a channel in the ER membrane) into the ER lumen. Once in the lumen, the signal peptide sequence itself is cleaved from the preproprotein. Subsequent biosynthesis occurs within the ER and Golgi, and it involves excision of peptides from the precursor by prohormone convertase enzymes, and the removal of remaining dibasic cleavage markers by carboxypeptidases. The resulting peptides are then available for posttranslational modifications, the most common of which is C-terminal amidation, which converts a C-terminal Glycine amino acid (found at the C-terminus of many encoded/immature neuropeptides) to amide (NH2). Amidation in flatworms is performed by the sequential actions of two enzymes, peptidylglycine α-hydroxylating monooxygenase (PHM) and peptidyl α-hydroxyglycine α-amidating lyase (PAL). In mammals, both of these functions are performed by a single bifunctional enzyme, peptidylglycine α-amidating monooxygenase (PAM). Amidated peptides are considered mature. They are stored in presynaptic secretory vesicles, from which they are exocytosed into the synapse when required.

Neuropeptide Classification

All flatworm neuropeptides are synthesized essentially by the mechanisms described earlier, but they display important differences in structure and function derived from primary amino acid sequence and the presence/absence of posttranslational modifications. Recent years have seen improved understanding of the sequence diversity of flatworm neuropeptide genes, thanks largely to improved genomic sequence resources. Prior to the genomic era, neuropeptide structure and sequence data were derived from HPLC-based isolation and peptide sequencing of peaks/fractions with immunoreactivity to known peptides from other species (Day & Maule, 1999). These methods yielded seven flatworm peptides—five FMRFamide-like peptides (FLPs) (Maule et al., 1993, 1994; Johnston et al., 1995, 1996) and two neuropeptide Fs (NPFs) (Maule et al., 1991; Curry et al., 1992). These isolations are particularly impressive given the technical difficulties associated with detecting peptides that are generally of low abundance from, as is the case with parasite-derived peptides, biological samples that are difficult to obtain in large quantities. For example, the extraction of GNFFRFamide from Moniezia expansa required one kilogram of tissue (Maule et al., 1993). Given these technical limitations, it is perhaps understandable why no peptides have been biochemically isolated from any trematode species. Subsequent work employed molecular techniques to clone the genes encoding some of these peptides in cestodes, trematodes, and planaria (Mair et al., 2000; Dougan et al., 2002; Humphries et al., 2004), but not until the arrival of the genomic era did we appreciate the full complexity of the flatworm neuropeptidergic system. Table 8.1 illustrates our current knowledge of neuropeptide precursor genes in phylum Platyhelminthes, based on published data derived from genome or transcriptome datasets. Sixty-four neuropeptide-encoding genes are known in the phylum, although these data are derived from only 12 species, and the distribution of homologues between species remains uncertain. It is unclear whether gaps in the interspecies distributions of npp homologues indicate a species-specific component of neuropeptide expression, or simply the incomplete nature of many of these datasets. S. mediterrannea represents the current most complete dataset, as reflected by its 51 precursors and 142 validated peptides (Collins et al., 2010). MS-validated peptides from this study were also queried against the genomes of S. japonicum and S. mansoni, allowing Collins et al. to identify an additional eight schistosome prepropeptide genes, including the YIRFamide encoding npp-23. YIRFamide was originally isolated from Bdelloura candida, a marine turbellarian, and is one of the most commonly tested peptides in flatworm bioassays (see later). Native Schmidtea peptides have not yet been searched against other flatworm datasets, so their conservation across the phylum remains unknown.

The flatworm neuropeptide dataset contains homologues of recognized invertebrate (p. 231) peptide families, including the FLP and NPF/NPY families, as well as APGWamide, buccalin, and myomodulin. S. mansoni also encodes peptides very similar to vertebrate neuropeptide FF (McVeigh et al., 2009). Schmidtea possesses additional homologues of cerebral peptide, gonadotropin-releasing hormone, insulin-like peptide, pyrokinin, and pedal peptide precursors (Collins et al., 2010). Based on the varied roles of their homologues in other invertebrates, it seems reasonable to posit that flatworm peptides fulfil a broad range of functions in addition to the more commonly ascribed neuromuscular roles of FLPs. Notably, with the exception of FLPs and NPFs, most of these peptide families have not yet been tested in flatworm neuromuscular bioassays.

Neuropeptide Signaling and Functions

Silencing components of the neuropeptide biosynthetic pathway illustrates the importance of peptidergic signaling for flatworms. RNAi of a PC2-like prohormone convertase in S. mediterrannea correlates with globally reduced levels of properly processed peptides in MS-analyzed tissue extracts. This intervention interferes with normal motility/behavior, regeneration, and reproductive development (Reddien et al., 2005; Collins et al., 2010). Similar phenotypes were reported following knockdown of a PC2 chaperone protein (7B2) and regulator of G protein signaling (RGS) proteins (Reddien et al., 2005), suggesting that neuropeptide biosynthesis and function are generally important for normal helminth behavior. PC2 knockdown has also been used to demonstrate a link between neuropeptide signaling and pharynx extension in planarians (Shimoyama et al., 2016).

FLP Structure and Function

FLPs (referred to in early literature as FMRFamide-related peptides, FaRPs), represent the largest known family of bioactive peptides in nature (McVeigh et al., 2006). Originally named based on sequence similarity with the eponymous FMRFamide, a cardioactive tetrapeptide in molluscs, FLPs are defined as short, bioactive amidated peptides with a C-terminal RFamide motif (McVeigh et al., 2009). Such peptides are now recognized as common throughout the protostomian invertebrate lineage. In deuterostomes the family has diversified to encompass several subfamilies bearing C-terminal RFamide motifs (e.g., gonadotropin inhibitory hormone, neuropeptide FF, prolactin-releasing peptide) (Elphick & Mirabeau, 2014). Flatworms possess at least seven FLP-encoding gene homologues (Table 8.1).

FLPs trigger a consistent excitatory/contractile response in whole-worm assays, muscle strip preparations, and dispersed muscle fibers from several species, with qualitatively similar excitatory responses also triggered by FLPs of nematode or arthropod origin (Mousley et al., 2004). The universally excitatory impact of FLPs suggests the existence of a monofunctional FLP receptor-signaling pathway in flatworms, data that are particularly intriguing given the multifunctional motility impacts of FLP families in other invertebrate genera (Mercier et al., 2003; R. Nichols, 2003; López-Vera et al., 2008). An important advance was the development of a muscle cell dispersal assay that enabled peptides to be tested on functional muscle cells that had been enzymatically digested and separated from other tissue (Blair et al., 1991). Significantly, this assay allowed experimentation in the absence of neuronal tissue; in the case of whole-worm and muscle strip assays, the relative impacts of test compounds on nerve and muscle cannot be discerned. Dispersed cell experiments illustrated the presence of a FLP-activated receptor on flatworm muscle tissue, implicating FLPs in the direct control of flatworm neuromuscular function (Day et al., 1994, 1997; Moneypenny et al., 2001).

FLPs in other invertebrates exert their functions primarily through GPCRs (Nichols, 2003; Claeys et al., 2005; Peymen et al., 2014; Zatylny-Gaudin & Favrel, 2014). A notable exception is the FMRFamide-activated Na+ channel (FaNaC), through which FMRFamide mediates fast neuronal transmission in gastropod molluscs and cnidarians (Lingueglia et al., 1995; Golubovic et al., 2007). There is also some electrophysiology evidence for a FLP-gated Cl channel in nematodes (Maule et al., 1995; Holden-Dye et al., 1997; Purcell et al., 2002). There is currently no evidence for peptide-gated ion channels in flatworms.

Only one FLP-activated flatworm GPCR has been directly identified, from the planarian D. tigrina (GtNPR1; Omar et al., 2007; Zamanian et al., 2012). In a heterologous expression system, this receptor responded most potently to GYIRFamide (Fig. 8.2). Although also activated by peptides encoded by the nematode flp-1 gene, GtNPR1 did not respond to flatworm NPFs. A subsequent RNAi-based deorphanization study confirmed GYIRFamide’s activation of GtNPR1 and showed that endogenous GtNPR1 signaled through a Gαi/o pathway (Zamanian et al., 2012; (p. 232) (p. 233) (p. 234) (p. 235) (p. 236) Fig. 8.2). Although these studies are promising demonstrations of the feasibility of flatworm peptide receptor deorphanization, further work is required before GtNPR1 or its parasite homologues can be considered as candidate anthelmintic drug targets. Notably, Zamanian et al. (2012) reported no phenotypic consequences of silencing GtNPR1, although it is unclear whether detailed phenotypic screens were performed in this study. In any case, the absence of obviously aberrant motility phenotypes suggests that GtNPR1 is probably not a muscle-based FLP receptor; GtNPR1 currently lacks tissue expression data that would either support or refute this hypothesis. Further studies are required to clarify both of these points. In terms of localization, we also lack any evidence that GYIRFamide and GtNPR1 are expressed in a manner that would allow their interaction in vivo. One major advantage of GtNPR1 as a target for compound screens is its clear amenability to heterologous expression in a high throughput screen (HTS)-compatible system. The argument for committing GtNPR1 to HTS of pharmaceutical compound libraries could be solidified by addressing the knowledge gaps pointed out earlier. These data would help ascertain GtNPR1’s candidacy by contributing to a “validation portfolio” of relevant biological data that would permit evaluation of the potential of this parasite receptor as a drug target candidate.

Table 8.1 Flatworm Neuropeptide Precursor Genes Identified from Sequence Datasets

Flatworm Neurobiology in the Postgenomic Era

Flatworm Neurobiology in the Postgenomic Era

Flatworm Neurobiology in the Postgenomic Era

Flatworm Neurobiology in the Postgenomic Era

Notes: Predicted peptide encoding genes are shown, alongside the peptide family encoded by each gene, as described by (a) McVeigh et al., 2009, (b) Berriman et al., 2009, (c) Collins et al., 2010, and (d) Tsai et al., 2013. Representation of each gene in genome or transcriptome datasets is identified by shading. Where more than one copy of a gene is present (i.e., two or more genes encoding essentially identical peptides), the number of copies is denoted by a number. Asterisks identify genes from which predicted peptides have been validated by mass spectrometry (Collins et al., 2010).

cpp, cerebral peptide prohormone; grh, gonadotropin releasing hormone-like; ilp, insulin-like peptide; mpl, myomodulin prohormone-like; npp, neuropeptide precursor; npy, neuropeptide Y superfamily; ppl, pyrokinin prohormone like; ppp, pedal peptide prohormone like; spp, secreted peptide prohormone.

In addition to GtNPR1, which signals through Gαi/o and adenylate cyclase (Zamanian et al., 2012), there is biochemical evidence for additional FLP-activated signaling pathways in trematodes. In F. hepatica, GYIRFamide activates a phospholipase C (PLC)/protein kinase C (PKC)–coupled receptor pathway in tissue homogenates (Graham et al., 2000). Given the crude nature of the whole-worm extracts used in these assays, it is impossible to ascertain from which tissues these receptor signals originated. Although employing very different methods, a comparison of Graham et al. (2000)’s results with those of Zamanian et al. (2012), discussed earlier, suggests that GYIRFamide operates through distinct GPCR signaling axes in trematodes (Gαi/o-adenylate cyclase) and planarians (Gαq-PLC). If such interspecies distinctions hold true for other receptors, we may find that planarians are of limited utility as models for parasitic forms.

Novozhilova et al. (2010) demonstrated that YIRFamide’s contractile effect on S. mansoni muscle fibers was mediated through a PKC-dependent pathway, triggering cytosolic Ca2+ influx through sarcolemmal voltage-operated Ca2+ channels (VOCCs).

There is appreciable evidence of a central role for FLPergic signaling in flatworm neuromuscular function, and these data frame flatworm FLP signaling as a source of potential therapeutic targets. We still lack many of the core tools required to exploit neuropeptide receptors and associated proteins for drug discovery, most notably the absence of any deorphanized parasite FLP receptors.

NPF/NPY Structure and Function

NPFs have been isolated biochemically from two flatworm species, the cestode Moniezia expansa and the turbellarian Arthurdendyus triangulatus (Maule et al., 1991; Curry et al., 1992). Attempts to isolate NPF from schistosomes were unsuccessful even when using 10,000 adult S. mansoni (Humphries et al., 2004). These isolations were stimulated by studies reporting immunoreactivity to members of the vertebrate neuropeptide Y superfamily in flatworm nervous systems, particularly to pancreatic polypeptide (PP). Chromatographic experiments showed that the single PP immunoreactive peaks in extracts from Diclidophora merlangi, M. expansa, and S. mansoni coeluted with other NPF/NPY peptides, suggesting that these peaks contained peptides with NPY superfamily-like sizes and hydrophobicities, and implying the presence of NPF-like peptides in these species (McVeigh et al., 2005b). Extraction of cestode and turbellarian NPFs, and subsequent cloning of their encoding genes, provided the first evidence of flatworm NPF structure. Flatworm NPFs are 29–47 amino acids in length, with conserved motifs consisting of a C-terminal GRPRFamide, with conserved tyrosines at positions 10 and 17 upstream of the C-terminus. Classically, NPFs (named for their C-terminal Phe) were considered the invertebrate version of NPY (named for C-terminal Tyr), which itself was considered restricted to vertebrates. Genomic data have now revealed that both NPF and NPY peptides can be present in individual flatworm species. Whether there are differences in function or signaling between these peptides remains unknown. Genomic structure provides further evidence for the relatedness of NPF/NPY, since almost all NPF and NPY encoding genes described to date have a conserved phase 2 intron in the codon for their penultimate Arg residue (Larhammar et al., 1987; Mair et al., (p. 237) 2000; Dougan et al., 2002; McVeigh et al., 2009; Collins et al., 2010).

NPY superfamily peptides signal exclusively through GPCRs. Various NPY receptor subfamilies are recognized, including Y1, Y2, Y4, Y5, and Y6 (Larhammar & Salaneck, 2004), each of which varies in its occurrence and complexity between vertebrate species. Invertebrate NPF receptors are also GPCRs, but they are characterized by lower gene complexity—only one or two NPF receptors are present in insect and mollusc genomes (Nässel & Wegener, 2011; Vogel et al., 2013). NPF is involved with functions relating particularly to feeding/appetite, as well as learning, stress, and alcohol tolerance in insects and molluscs (Nässel & Wegener, 2011). While no flatworm NPF receptors have been cloned, available evidence suggests that NPFs trigger conserved NPY-like signaling pathways. NPY signals through Gαi/o by inhibiting the accumulation of cAMP (Motulsky & Michel, 1988; Aakerlund et al., 1990). Similarly in schistosome homogenates, NPF inhibits the accumulation of cAMP at concentrations as low as 10–11 M (Humphries et al., 2004).

Data on the functions of flatworm NPFs are similarly sparse. There is some evidence from S. mediterranea to implicate an NPF (Smed-npy-8), and its receptor, in reproductive function (Collins et al., 2010; Saberi et al., 2016). Knockdown of these genes led to phenotypes indicating a loss of sexual maturity, including regressed testes, cessation of sperm production, regression of copulatory organs, and in some cases, the disappearance of the gonopore. These physical phenotypes correlated with interrupted expression of developmental genes in affected tissues, suggesting that NPY-8 is essential for the normal maintenance of reproductive maturity that is required for planarian reproduction. NPF may also have a role in the control of cellular regeneration in planarians, given that exogenously applied NPF stimulates the mitotic activity of neoblasts, which translates to accelerated regeneration of the head, nervous system, and musculature (Kreshchenko et al., 2008).


Peptides with L/M/Iamide C-termini comprise the largest neuropeptide family in flatworms, with such peptides encoded by 21 of the 64 known neuropeptide genes (Berriman et al., 2009; McVeigh et al., 2009; Collins et al., 2010; Tsai et al., 2013). Some known peptides bear this motif, including molluscan buccalin (Aplysia: GMDSLAFSGGLamide; Cropper et al., 1988), myomodulin (Aplysia: PMSMLRLamide; Cropper et al., 1987), and arthropod pyrokinin (Leucophora: GTSPTPRLamide; Holman et al., 1986) neuropeptides. Buccalin, myomodulin, and pyrokinin are all myoexcitatory in their species of origin, so we would hypothesize that their flatworm homologues are similarly myoactive. Only one study has examined the bioactivity of a native flatworm member of this family. GFVRIamide, one of the peptides produced by the npp-1 gene, localizes to two neurons in the schistosome CNS and has inhibitory impacts on motility following application to whole-adult S. mansoni (McVeigh et al., 2011). Attempts to silence Sm-npp-1 were uninformative; although npp-1 transcripts were potently and specifically knocked down in schistosomulae, no aberrant motility phenotypes were noted in these worms. Arthropod type-A allatostatins also bear C-terminal L/Iamide signatures (characteristic motif: (Y/F)XFG(L/I)amide) and have myoactive effects in their host species. Allatostatins also trigger contractions of flatworm dispersed muscle fibers, in an example of interphyla functionality (Mousley et al., 2005).

Other Neuropeptides

Several additional smaller neuropeptide families have also been described from flatworms (McVeigh et al., 2009; Collins et al., 2010), for which limited data are available. These include the PWamides encoded by npp-5, which have been localized immunocytochemically to the S. mansoni PNS, including in innervation of muscle and reproductive organs (McVeigh et al., 2009). Although no somatostatin-like peptides have been identified, there is some evidence for the presence of somatostatin receptor-like GPCRs on the S. mansoni tegument (Chatterjee et al., 2005, 2007).


The neurobiology of parasitic flatworms appears to offer many opportunities for chemotherapeutic targeting in pathogenic helminths. The rapid development of genomics/transcriptomics, functional genomics, and heterologous expression methods during the last decade have enabled helminth parasitology to move from studying putative therapeutic targets based almost entirely on biochemistry, localization, and/or neuromuscular physiology toward the current paradigm that aims to apply systems biology methods to the identification, validation, and exploitation of potential chemotherapeutic targets. (p. 238) As detailed herein, these approaches have identified a cohort of heterologously expressed receptors, ion channels, and reuptake pumps, a handful of which have additionally been functionally validated by RNAi. These data, taken alongside improved knowledge of neuropeptides and biosynthetic enzyme complements, represent tangible progress in flatworm neurobiology during the past decade. Nevertheless, much remains to be done if this promise is to be translated into concrete progress with respect to developing new therapeutic options with which to treat helminth infections.

The receptors, transporters, and ion channels discussed earlier represent eminently druggable proteins, for which multiple classes of active compounds already exist. In addition to this appreciable pharmacopeia, it is particularly noteworthy that all of the proteins detailed earlier as successfully expressed in heterologous systems are described as “pharmacologically distinct” from their mammalian homologues. This implies considerable promise in the potential to target these proteins in a selective manner that would avoid nonspecific impacts on the host.


Aakerlund, L., Gether, U., Fuhlendorff, J., Schwartz, T. W., & Thastrup, O. (1990). Y1 receptors for neuropeptide Y are coupled to mobilization of intracellular calcium and inhibition of adenylate cyclase. FEBS Letters, 260(1), 73–78. this resource:

Agboh, K. C., Webb, T. E., Evans, R. J., & Ennion, S. J. (2004). Functional characterization of a P2X receptor from Schistosoma mansoni. The Journal of Biological Chemistry, 279(40), 41650–41657. doi:10.1074/jbc.M408203200.Find this resource:

Baxter, D. A., & Byrne, J. H. (2006). Feeding behavior of Aplysia: A model system for comparing cellular mechanisms of classical and operant conditioning. Learning & Memory, 13(6), 669–680. doi:10.1101/lm.339206.Find this resource:

Beaulieu, J.-M., & Gainetdinov, R. R. (2011). The physiology, signaling, and pharmacology of dopamine receptors. Pharmacological Reviews, 63(1), 182–217. doi:10.1124 /pr.110.002642.Find this resource:

Beech, R. N., Callanan, M. K., Rao, V. T. S., Dawe, G. B., & Forrester, S. G. (2013). Characterization of Cys-Loop receptor genes involved in inhibitory amine neurotransmission in parasitic and free living nematodes. Parasitology International, 62(6), 599–605. doi:10.1016/j.parint.2013.03.010.Find this resource:

Bentley, G. N., Jones, A. K., & Agnew, A. (2007). ShAR2β, a divergent nicotinic acetylcholine receptor subunit from the blood fluke schistosoma. Parasitology, 134(6), 833. doi:10.1017/S0031182006002162.Find this resource:

Bentley, G. N., Jones, A. K., & Agnew, A. (2003). Mapping and sequencing of acetylcholinesterase genes from the platyhelminth blood fluke Schistosoma. Gene, 314 (September), 103–112. this resource:

Bentley, G. N., Jones, A. K., & Agnew, A. (2005). Expression and comparative functional characterisation of recombinant acetylcholinesterase from three species of Schistosoma. Molecular and Biochemical Parasitology, 141(1), 119–123. doi:10.1016/j.molbiopara.2005.01.019.Find this resource:

Bentley, G. N., Jones, A. K., Oliveros Parra, W. G., & Agnew, A. (2004). ShAR1alpha and ShAR1beta: Novel putative nicotinic acetylcholine receptor subunits from the platyhelminth blood fluke Schistosoma. Gene, 329(March), 27–38. doi:10.1016/j.gene.2003.12.009.Find this resource:

Berriman, M., Haas, B. J., LoVerde, P. T., Wilson, R. A., Dillon, G. P., Cerqueira, G. C., . . . El-Sayed, N. M. (2009). The genome of the blood fluke Schistosoma mansoni. Nature, 460(7253), 352–358. this resource:

Bian, K., & Murad, F. (2014). What is next in nitric oxide research? From cardiovascular system to cancer biology. Nitric Oxide: Biology and Chemistry/Official Journal of the Nitric Oxide Society, 43(December), 3–7. doi:10.1016 /j.niox.2014.08.006.Find this resource:

Blair, K. L., & Anderson, P. A. V. (2009). Physiological and pharmacological properties of muscle cells isolated from the flatworm Bdelloura candida (Tricladia). Parasitology, 109(3), 325–335. doi:10.1017/S0031182000078355.Find this resource:

Blair, K. L., Day, T. A., Lewis, M. C., Bennett, J. L., & Pax, R. A. (1991). Studies on muscle cells isolated from Schistosoma mansoni: A Ca(2+)-dependent K+ channel. Parasitology, 102 Pt 2 (April), 251–258. this resource:

Blenau, W., & Thamm, M. (2011). Blenau distribution of serotonin (5-HT) and its receptors in the insect brain with focus on the mushroom bodies: Lessons from Drosophila melanogaster and Apis mellifera. Arthropod Structure and Development, 40(5), 381–394.Find this resource:

Boyle, J. P., Hillyer, J. F., & Yoshino, T. P. (2003). Pharmacological and autoradiographical characterization of serotonin transporter-like activity in sporocysts of the human blood fluke, Schistosoma mansoni. Journal of Comparative Physiology. A, Neuroethology, Sensory, Neural, and Behavioral Physiology, 189(8), 631–641. doi:10.1007/s00359-003-0429-8.Find this resource:

Brockie, P. (2006). Ionotropic glutamate receptors: Genetics, behavior and electrophysiology. WormBook, January, 1–16. doi:10.1895/wormbook.1.61.1.Find this resource:

Brown, K. M., Roy, K. K., Hockerman, G. H., Doerksen, R. J., & Colby, D. A. (2015). Activation of the γ-aminobutyric acid type B (GABA(B)) receptor by agonists and positive allosteric modulators. Journal of Medicinal Chemistry, 58(16), 6336–6347. doi:10.1021/jm5018913.Find this resource:

Burnstock, G. (2013). Glioma signaling. Edited by Jolanta Barańska. Advances in Experimental Medicine and Biology. Vol. 986. Dordrecht, the Netherlands: Springer. doi:10.1007 /978-94-007-4719-7.Find this resource:

Camacho, M., & Agnew, A. (1995). Schistosoma: Rate of glucose import is altered by acetylcholine interaction with tegumental acetylcholine receptors and acetylcholinesterase. Experimental Parasitology, 81(4), 584–591. doi:10.1006 /expr.1995.1152.Find this resource:

Camicia, F., Herz, M., Prada, L. C., Kamenetzky, L., Simonetta, S. H., Cucher, M. A., . . . Rosenzvit, M. C. (2013). The nervous and prenervous roles of serotonin in Echinococcus spp. International Journal for Parasitology, 43(8), 647–659. doi:10.1016/j.ijpara.2013.03.006.Find this resource:

Campos, T. D. L., Young, N. D., Korhonen, P. K., Hall, R. S., Mangiola, S., Lonie, A., & Gasser, R. B. (2014). Identification of G protein-coupled receptors in Schistosoma haematobium (p. 239) and S. mansoni by comparative genomics. Parasites & Vectors, 7(1), 242. doi:10.1186/1756-3305-7-242.Find this resource:

Caveney, S., Cladman, W., Verellen, L., & Donly, C. (2006). Ancestry of neuronal monoamine transporters in the metazoa. The Journal of Experimental Biology, 209(Pt 24), 4858–4868. doi:10.1242/jeb.02607.Find this resource:

Chatterjee, S., De Beeck, J. O., Rao, A. V., Desai, D. V., Vrolix, G., Rylant, F., . . . Van Marck, E. (2007). Prolonged somatostatin therapy may cause down-regulation of SSTR-like GPCRs on Schistosoma mansoni. Journal of Vector Borne Diseases, 44(3), 164–180. this resource:

Chatterjee, S., Vrolix, G., Depoortere, I., Peeters, T., & Van Marck, E. (2005). The therapeutic effect of the neuropeptide hormone somatostatin on Schistosoma mansoni caused liver fibrosis. BMC Infectious Diseases, 5(January), 45. doi:10.1186/1471-2334-5-45.Find this resource:

Claeys, I., Poels, J., Simonet, G., Franssens, V., Van Loy, T., Van Hiel, M. B., . . . Broeck, J. V. (2005). Insect neuropeptide and peptide hormone receptors: Current knowledge and future directions. Vitamins and Hormones, 73(January), 217–282. doi:10.1016/S0083-6729(05)73007-7.Find this resource:

Collingridge, G. L., Olsen, R. W., Peters, J., & Spedding, M. (2009). A nomenclature for ligand-gated ion channels. Neuropharmacology, 56(1), 2–5. doi:10.1016 /j.neuropharm.2008.06.063.Find this resource:

Collins, J. J., Hou, X., Romanova, E. V., Lambrus, B. G., Miller, C. M., Saberi, A., . . . Newmark, P. A. (2010). Genome-wide analyses reveal a role for peptide hormones in planarian germline development. PLoS Biology, 8(10), e1000509. doi:10.1371/journal.pbio.1000509.Find this resource:

Cropper, E. C., Miller, M. W., Tenenbaum, R., Kolks, M. A., Kupfermann, I., & Weiss, K. R. (1988). Structure and action of buccalin: A nodulatory neuropeptide localized to an identified small cardioactive peptide-containing cholinergic motor neuron of Aplysia californica. Proceedings of the National Academy of Sciences of the United States of America, 85(16), 6177–6181. this resource:

Cropper, E. C., Tenenbaum, R., Kolks, M. A., Kupfermann, I., & Weiss, K. R. (1987). Myomodulin: A bioactive neuropeptide present in an identified cholinergic buccal motor neuron of Aplysia. Proceedings of the National Academy of Sciences of the United States of America, 84(15), 5483–5486. this resource:

Curry, W. J., Shaw, C., Johnston, C. F., Thim, L., & Buchanan, K. D. (1992). Neuropeptide F: Primary structure from the tubellarian, Artioposthia triangulata. Comparative Biochemistry and Physiology. C, Comparative Pharmacology and Toxicology, 101(2), 269–274. this resource:

Da’dara, A., & Skelly, P. J. (2011). Manipulation of vascular function by blood flukes? Blood Reviews, 25(4), 175–179. doi:10.1016/j.blre.2011.04.002.Find this resource:

Davis, R., & Stretton, A. O. W. (1995). Neurotransmitters of helminths. In J. Marr & M. Muller (Eds.), Biochemistry and molecular biology of parasites (pp. 257–287). London: Academic Press.Find this resource:

Day, T. A., Bennett, J. L., & Pax, R. A. (1994). Serotonin and its requirement for maintenance of contractility in muscle fibres isolated from Schistosoma mansoni. Parasitology, 108(Pt 4, May), 425–432. this resource:

Day, T. A., Chen, G. Z., Miller, C., Tian, M., Bennett, J. L., & Pax, R. A. (1996). Cholinergic inhibition of muscle fibres isolated from Schistosoma mansoni (Trematoda:Digenea). Parasitology, 113(Pt 1 July), 55–61. this resource:

Day, T. A., & Maule, A. G. (1999). Parasitic peptides! The structure and function of neuropeptides in parasitic worms. Peptides, 20(8), 999–1019. doi:10.1016 /S0196-9781(99)00093-5.Find this resource:

Day, T. A., Maule, A. G., Shaw, C., Halton, D. W., Moore, S., Bennett, J. L., & Pax, R. L. (1994). Platyhelminth FMRFamide-related peptides (FaRPs) contract Schistosoma mansoni (Trematoda: Digenea) muscle fibres in vitro. Parasitology, 109(Pt 4, November), 455–459. this resource:

Day, T. A., Maule, A. G., Shaw, C., & Pax, R. A. (1997). Structure-activity relationships of FMRFamide-related peptides contracting Schistosoma mansoni muscle. Peptides, 18(7), 917–921. this resource:

Dernovici, S., Starc, T., Dent, J. A., & Ribeiro, P. (2007). The serotonin receptor SER-1 (5HT2ce) contributes to the regulation of locomotion in Caenorhabditis elegans. Developmental Neurobiology, 67(2), 189–204. doi:10.1002/dneu.20340.Find this resource:

Dougan, P. M., Mair, G. R., Halton, D. W., Curry, W. J., Day, T. A., & Maule, A. G. (2002). Gene organization and expression of a neuropeptide Y homolog from the land planarian Arthurdendyus triangulatus. The Journal of Comparative Neurology, 454(1), 58–64. doi:10.1002/cne.10440.Find this resource:

Dufour, V., Beech, R. N., Wever, C., Dent, J. A., & Geary, T. G. (2013). Molecular cloning and characterization of novel glutamate-gated chloride channel subunits from Schistosoma mansoni. PLoS Pathogens, 9(8), e1003586. doi:10.1371 /journal.ppat.1003586.Find this resource:

El-Shehabi, F., & Ribeiro, P. (2010). Histamine signalling in Schistosoma mansoni: Immunolocalisation and characterisation of a new histamine-responsive receptor (SmGPR-2). International Journal for Parasitology, 40(12), 1395–1406. doi:10.1016/j.ijpara.2010.04.006.Find this resource:

El-Shehabi, F., Taman, A., Moali, L. S., El-Sakkary, N., & Ribeiro, P. (2012). A novel G protein-coupled receptor of Schistosoma mansoni (SmGPR-3) is activated by dopamine and is widely expressed in the nervous system. PLoS Neglected Tropical Diseases, 6(2), e1523. doi:10.1371 /journal.pntd.0001523.Find this resource:

El-Shehabi, F., Vermeire, J. J., Yoshino, T. P., & Ribeiro, P. (2009). Developmental expression analysis and immunolocalization of a biogenic amine receptor in Schistosoma mansoni. Experimental Parasitology, 122(1), 17–27. doi:10.1016 /j.exppara.2009.01.001.Find this resource:

Elphick, M. R., & Mirabeau, O. (2014). The evolution and variety of RFamide-type neuropeptides: Insights from Deuterostomian invertebrates. Frontiers in Endocrinology, 5(January), 93. doi:10.3389/fendo.2014.00093.Find this resource:

Farooqui, T. (2007). Octopamine-mediated neuromodulation of insect senses. Neurochemical Research, 32(9), 1511–1529. doi:10.1007/s11064-007-9344-7.Find this resource:

Fontana, A. C. K., Sonders, M. S., Pereira-Junior, O. S., Knight, M., Javitch, J. A., Rodrigues, V., . . . Mortensen, O. V. (2009). Two allelic isoforms of the serotonin transporter from Schistosoma mansoni display electrogenic transport and high (p. 240) selectivity for serotonin. European Journal of Pharmacology, 616(1–3), 48–57. doi:10.1016/j.ejphar.2009.06.023.Find this resource:

Geary, T. G. (2010). Nonpeptide ligands for peptidergic G protein-coupled receptors. Advances in Experimental Medicine and Biology, 692(January), 10–26. this resource:

Geary, T. G., Marks, N. G., Maule, A. G., Bowman, J. W., Alexander-Bowman, S. J., Day, T. A., . . .Thompson, D. P. (1999). Pharmacology of FMRFamide-related peptides in helminths. Annals of the New York Academy of Sciences, 897(January), 212–227. this resource:

Geary, T. G., Sakanari, J. A., & Caffrey, C. R. (2015). Anthelmintic drug discovery: Into the future. The Journal of Parasitology 101 (2): 125–133. doi:10.1645/14-703.1.Find this resource:

Gether, U., Andersen, P. H., Larsson, O. M., & Schousboe, A. (2006). Neurotransmitter transporters: Molecular function of important drug targets. Trends in Pharmacological Sciences, 27(7), 375–383. doi:10.1016/ this resource:

Golubovic, A., Kuhn, A., Williamson, M., Kalbacher, H., Holstein, T. W., Grimmelikhuijzen, C. J. P., & Gründer, S. (2007). A peptide-gated ion channel from the freshwater polyp hydra. The Journal of Biological Chemistry, 282(48), 35098–35103. doi:10.1074/jbc.M706849200.Find this resource:

Graft, D. J., & Read, C. P. (1967). Specific acetylcholinesterase in Hymenolepis diminuta. The Journal of Parasitology, 53(5), 1030–1031. this resource:

Graham, M. K., Fairweather, I., & McGeown, J. G. (2000). Second messengers mediating mechanical responses to the FARP GYIRFamide in the fluke Fasciola hepatica. American Journal of Physiology. Regulatory, Integrative and Comparative Physiology, 279(6), R2089–R2094. this resource:

Halton, D. W. (1967). Histochemical studies of carboxylic esterase activity in Fasciola hepatica. The Journal of Parasitology, 53(6), 1210–1216. this resource:

Halton, D. W., & Gustafsson, M. K. S. (1996). Functional morphology of the platyhelminth nervous system. Parasitology, 113, S47–72.Find this resource:

Halton, D. W., & Maule, A. G. (2004). Flatworm nerve-muscle: Structural and functional analysis. Canadian Journal of Zoology, 82, 316–333.Find this resource:

Hamdan, F. F., Abramovitz, M., Mousa, A., Xie, J., Durocher, Y., & Ribeiro, P. (2002). A novel Schistosoma mansoni G protein-coupled receptor is responsive to histamine. Molecular and Biochemical Parasitology, 119(1), 75–86. this resource:

Hamdan, F. F., & Ribeiro, P. (1998). Cloning and characterization of a novel form of tyrosine hydroxylase from the human parasite, Schistosoma mansoni. Journal of Neurochemistry, 71(4), 1369–1380. this resource:

Hamdan, F. F., & Ribeiro, P. (1999). Characterization of a stable form of tryptophan hydroxylase from the human parasite Schistosoma mansoni. The Journal of Biological Chemistry, 274(31), 21746–21754. this resource:

Hillman, G. R., & Senft, A. W. (1973). Schistosome motility measurements: Response to drugs. The Journal of Pharmacology and Experimental Therapeutics, 185(2), 177–184. this resource:

Holden-Dye, L., Brownlee, D. J., & Walker, R. J. (1997). The effects of the peptide KPNFIRFamide (PF4) on the somatic muscle cells of the parasitic nematode Ascaris suum. British Journal of Pharmacology, 120(3), 379–386. doi:10.1038 /sj.bjp.0700906.Find this resource:

Holman, G. M., Cook, B. J., & Nachman, R. J. (1986). Primary structure and synthesis of a blocked myotropic neuropeptide isolated from the cockroach, Leucophaea maderae. Comparative Biochemistry and Physiology. C, Comparative Pharmacology and Toxicology, 85(1), 219–224. this resource:

Holmes, S. D., & Fairweather, I. (1984). Fasciola hepatica: The effects of neuropharmacological agents upon in vitro motility. Experimental Parasitology, 58(2), 194–208. this resource:

Hong, W. J., & Lev, S. (2014). Tethering the assembly of SNARE complexes. Trends in Cell Biology, 24(1), 35–43. doi:10.1016/j.tcb.2013.09.006.Find this resource:

Hope, B. T., Michael, G. J., Knigge, K. M., & Vincent, S. R. (1991). Neuronal NADPH diaphorase is a nitric oxide synthase. Proceedings of the National Academy of Sciences of the United States of America, 88(7), 2811–2814. this resource:

Hu, Y., Shi, D., Luo, Q., Liu, Q., Zhou, Y., Liu, L., . . . Shen, J. (2011). Cloning and characterization of a novel enzyme: Tyrosine hydroxylase from Schistosoma japonicum. Parasitology Research, 109(4), 1065–1074. doi:10.1007/s00436-011-2347-y.Find this resource:

Humphries, J. E., Kimber, M. J., Barton, Y.-W., Hsu, W., Marks, N. J., Greer, B., . . . Day, T. A. (2004). Structure and bioactivity of Neuropeptide F from the human parasites Schistosoma mansoni and Schistosoma japonicum. Journal of Biological Chemistry, 279(38), 39880–39885. doi:10.1074 /jbc.M405624200.Find this resource:

Husson, S. J., & Schoofs, L. (2007a). Altered neuropeptide profile of Caenorhabditis elegans lacking the chaperone protein 7B2 as analyzed by mass spectrometry. FEBS Letters, 581(22), 4288–4292. doi:10.1016/j.febslet.2007.08.003.Find this resource:

Husson, S. J., & Schoofs, L. (2007b). Processing of neuropeptide precursors in Caenorhabditis elegans. Communications in Agricultural and Applied Biological Sciences, 72(1), 199–203. this resource:

Ignarro, L. J. (1990). Nitric oxide: A novel signal transduction mechanism for transcellular communication. Hypertension, 16(5), 477–483. this resource:

Johnston, R. N., Shaw, C., Halton, D. W., Verhaert, P., & Baguna, J. (1995). GYIRFamide: A novel FMRFAmide-related peptide (FaRP) from the triclad turbellarian, Dugesia tigrina. Biochemical and Biophysical Research Communications, 209(2), 689–697. doi:10.1006/bbrc.1995.1554.Find this resource:

Johnston, R. N., Shaw, C., Halton, D. W., Verhaert, P., Blair, K. L., Brennan, G. P., . . . Anderson, P. A. (1996). Isolation, localization, and bioactivity of the FMRFamide-related neuropeptides GYIRFamide and YIRFamide from the marine turbellarian Bdelloura candida. Journal of Neurochemistry, 67(2), 814–821. this resource:

Kammermeier, P. J., & Yun, J. (2005). Activation of metabotropic glutamate receptor 1 dimers requires glutamate binding in both subunits. The Journal of Pharmacology and Experimental Therapeutics, 312(2), 502–508. doi:10.1124 /jpet.104.073155.Find this resource:

(p. 241) Kniazeff, J., Bessis, A.-S., Maurel, D., Ansanay, H., Prézeau, L., & Pin, J.-P. (2004). Closed state of both binding domains of homodimeric mGlu receptors is required for full activity. Nature Structural & Molecular Biology, 11(8), 706–713. doi:10.1038/nsmb794.Find this resource:

Kobayashi, K. (2001). Role of catecholamine signaling in brain and nervous system functions: New insights from mouse molecular genetic study. The Journal of Investigative Dermatology. Symposium Proceedings/the Society for Investigative Dermatology, Inc. [and] European Society for Dermatological Research, 6(1), 115–121. doi:10.1046/j.0022-202x.2001.00011.x.Find this resource:

Kohn, A. B., Moroz, L. L., Lea, J. M., & Greenberg, R. M. (2001). Distribution of nitric oxide synthase immunoreactivity in the nervous system and peripheral tissues of Schistosoma mansoni. Parasitology, 122 Pt 1(January), 87–92. this resource:

Kreshchenko, N. D., Sedelnikov, Z., Sheiman, I. M., Reuter, M., Maule, A. G., & Gustafsson, M. K. S. (2008). Effects of Neuropeptide F on regeneration in Girardia tigrina (Platyhelminthes). Cell and Tissue Research, 331(3), 739–750. doi:10.1007/s00441-007-0519-y.Find this resource:

Lange, A. B. (2009). Tyramine: From octopamine precursor to neuroactive chemical in insects. General and Comparative Endocrinology, 162(1), 18–26. doi:10.1016 /j.ygcen.2008.05.021.Find this resource:

Larhammar, D., Ericsson, A., & Persson, H. (1987). Structure and expression of the rat neuropeptide Y gene. Proceedings of the National Academy of Sciences of the United States of America, 84(7), 2068–2072. this resource:

Larhammar, D., & Salaneck, E. (2004). Molecular evolution of NPY receptor subtypes. Neuropeptides, 38(4), 141–151. doi:10.1016/j.npep.2004.06.002.Find this resource:

Larsen, M. B., Fontana, A. C. K., Magalhães, L. G., Rodrigues, V., & Mortensen, O. V. (2011). A catecholamine transporter from the human parasite Schistosoma mansoni with low affinity for psychostimulants. Molecular and Biochemical Parasitology, 177(1), 35–41. doi:10.1016/j.molbiopara.2011.01.006.Find this resource:

Li, C., & Kim, K. (2008). Neuropeptides. WormBook: The Online Review of C. Elegans Biology, January, 1–36. doi:10.1895 /wormbook.1.142.1.Find this resource:

Lingueglia, E., Champigny, G., Lazdunski, M., & Barbry, P. (1995). Cloning of the amiloride-sensitive FMRFamide peptide-gated sodium channel. Nature, 378 (6558), 730–733. doi:10.1038/378730a0.Find this resource:

López-Vera, E., Aguilar, M. B., & de la Cotera, E. P. H. (2008). FMRFamide and related peptides in the phylum Mollusca. Peptides, 29(2), 310–317. doi:10.1016 /j.peptides.2007.09.025.Find this resource:

MacDonald, K., Buxton, S., Kimber, M. J., Day, T. A., Robertson, A. P., & Ribeiro, P. (2014). Functional characterization of a novel family of acetylcholine-gated chloride channels in Schistosoma mansoni. Edited by Robert M. Greenberg. PLoS Pathogens, 10(6), e1004181. doi:10.1371 /journal.ppat.1004181.Find this resource:

MacDonald, K., Kimber, M. J., Day, T. A., & Ribeiro, P. (2015). A constitutively active G protein-coupled acetylcholine receptor regulates motility of arval Schistosoma mansoni. Molecular and Biochemical Parasitology, 202(1), 29–37. doi:10.1016/j.molbiopara.2015.09.001.Find this resource:

Mair, G. R., Halton, D. W., Shaw, C., & Maule, A. G. (2000). The neuropeptide F (NPF) encoding gene from the cestode, Moniezia expansa. Parasitology, 120(Pt 1, January), 71–77. this resource:

Marks, N. J., & Maule, A. G. (2010). Neuropeptides in helminths: Occurrence and distribution. Advances in Experimental Medicine and Biology, 692(January), 49–77. this resource:

Maule, A. G., Halton, D. W., Allen, J. M., & Fairweather, I. (2009). Studies on motility in vitro of an ectoparasitic monogenean, Diclidophora merlangi. Parasitology, 98(1), 85–93. doi:10.1017/S0031182000059722.Find this resource:

Maule, A. G., Shaw, C., Bowman, J. W., Halton, D. W., Thompson, D. P., Thim, L., . . . Geary, T. G. (1995). Isolation and preliminary biological characterization of KPNFIRFamide, a novel FMRFamide-related peptide from the free-living nematode, Panagrellus redivivus. Peptides, 16(1), 87–93. this resource:

Maule, A. G., Shaw, C., Halton, D. W., Curry, W. J., & Thim, L. (1994). RYIRFamide: A turbellarian FMRFamide-related peptide (FaRP). Regulatory Peptides, 50(1), 37–43. this resource:

Maule, A. G., Mousley, A., Marks, N. J., Day, T. A., Thompson, D. P., Geary, T. G., & Halton, D. W. (2002). Neuropeptide signaling systems—Potential drug targets for parasite and pest control. Current Topics in Medicinal Chemistry, 2(7), 733–758. this resource:

Maule, A., Shaw, C., Halton, D., & Thim, L. (1993). GNFFRFamide: A novel FMRFamide-immunoreactive peptide isolated from the sheep tapeworm, Moniezia expansa. Biochemical and Biophysical Research Communications, 193 (3), 1054–1060. doi:10.1006/bbrc.1993.1732.Find this resource:

Maule, A. G., Shaw, C., Halton, D. W., Thim, L., Johnston, C. F., Fairweather, I., & Buchanan, K. D. (1991). Neuropeptide F: A novel parasitic flatworm regulatory peptide from Moniezia expansa (Cestoda: Cyclophyllidea). Parasitology, 102, 309–316.Find this resource:

McKay, D. M., Halton, D. W., Allen, J. M., & Fairweather, I. (1989). The effects of cholinergic and serotoninergic drugs on motility in vitro of Haplometra cylindracea (Trematoda: Digenea). Parasitology, 99, Pt 2 (October), 241–252. this resource:

McVeigh, P., Atkinson, L., Marks, N. J., Mousley, A., Dalzell, J. J., Sluder, A., . . . Maule, A. G. (2012). Parasite neuropeptide biology: Seeding rational drug target selection? International Journal for Parasitology. Drugs and Drug Resistance, 2(December), 76–91. doi:10.1016/j.ijpddr.2011.10.004.Find this resource:

McVeigh, P., Geary, T. G., Marks, N. J., & Maule, A. G. (2006). The FLP-side of nematodes. Trends in Parasitology, 22(8), 385–396. doi:10.1016/ this resource:

McVeigh, P., Kimber, M. J., Novozhilova, E., & Day, T. A. (2005b). Neuropeptide signalling systems in flatworms. Parasitology, 131, Suppl (January), S41–S55. doi:10.1017 /S0031182005008851.Find this resource:

McVeigh, P., Leech, S., Mair, G. R., Marks, N. J., Geary, T. G., & Maule, A. G. (2005a). Analysis of FMRFamide-like peptide (FLP) diversity in phylum Nematoda. International Journal for Parasitology, 35(10), 1043–1060. doi:10.1016/j.ijpara.2005.05.010.Find this resource:

McVeigh, P., Mair, G. R., Atkinson, L., Ladurner, P., Zamanian, M., Novozhilova, E., . . . Maule, A. G. (2009). Discovery of multiple neuropeptide families in the phylum platyhelminthes. International Journal for Parasitology, 39(11), 1243–1252. doi:10.1016/j.ijpara.2009.03.005.Find this resource:

(p. 242) McVeigh, P., Mair, G. R., Novozhilova, E., Day, A., Zamanian, M., Marks, N. J., . . . Maule, A. G. (2011). Schistosome I/lamides—A new family of bioactive helminth neuropeptides. International Journal for Parasitology, 41(8), 905–913. doi:10.1016/j.ijpara.2011.03.010.Find this resource:

Mellin, T. N., Busch, R. D., Wang, C. C., & Kath, G. (1983). Neuropharmacology of the parasitic trematode, Schistosoma mansoni. The American Journal of Tropical Medicine and Hygiene, 32(1), 83–93. this resource:

Mendonça-Silva, D. L., Gardino, P. F., Kubrusly, R. C. C., De Mello, F. G., & Noël, F. (2004). Characterization of a GABAergic neurotransmission in adult Schistosoma mansoni. Parasitology, 129(Pt 2), 137–146. this resource:

Mendonça-Silva, D. L., Pessôa, R. F., & Noël, F. (2002). Evidence for the presence of glutamatergic receptors in adult Schistosoma mansoni. Biochemical Pharmacology, 64(9), 1337–1344. this resource:

Mercier, A. J., Friedrich, R., & Boldt, M. (2003). Physiological functions of FMRFamide-like peptides (FLPs) in crustaceans. Microscopy Research and Technique, 60(3), 313–324. doi:10.1002/jemt.10270.Find this resource:

Messerli, S. M., Morgan, W., Birkeland, S. R., Bernier, J., Cipriano, M. J., McArthur, A. G., & Greenberg, R. M. (2006). Nitric oxide-dependent changes in Schistosoma mansoni gene expression. Molecular and Biochemical Parasitology, 150(2), 367–370. doi:10.1016/j.molbiopara.2006.08.003.Find this resource:

Miller, C. L., Day, T. A., Bennett, J. L., & Pax, R. A. (1996). Schistosoma mansoni: L-glutamate-induced contractions in isolated muscle fibers: Evidence for a glutamate transporter. Experimental Parasitology, 84(3), 410–419. doi:10.1006 /expr.1996.0129.Find this resource:

Moneypenny, C. G., Kreshchenko, N., Moffett, C. L., Halton, D. W., Day, T. A., & Maule, A. G. (2001). Physiological effects of FMRFamide-related peptides and classical transmitters on dispersed muscle fibres of the turbellarian, Procerodes littoralis. Parasitology, 122(Pt 4), 447–455. this resource:

Motulsky, H. J., & Michel, M. C. (1988). Neuropeptide Y mobilizes Ca2+ and inhibits adenylate cyclase in human erythroleukemia cells. The American Journal of Physiology, 255(6 Pt 1), E880–E885. this resource:

Mousley, A., Marks, N. J., & Maule, A. G. (2004). Neuropeptide signalling: A repository of targets for novel endectocides? Trends in Parasitology, 20(10), 482–487. doi:10.1016 / this resource:

Mousley, A., Maule, A. G., Halton, D. W., & Marks, N. J. (2005). Inter-phyla studies on neuropeptides: The potential for broad-spectrum anthelmintic and/or endectocide discovery. Parasitology, 131(Suppl. January), S143–S167. doi:10.1017/S0031182005008553.Find this resource:

Mousley, A., Moffett, C. L., Duve, H., Thorpe, A., Halton, D. W., Geary, T. G., . . . Marks, N. J. (2005). Expression and bioactivity of allatostatin-like neuropeptides in helminths. International Journal for Parasitology, 35(14), 1557–1567. doi:10.1016/j.ijpara.2005.08.002.Find this resource:

Mutafova-Yambolieva, V. N., & Durnin, L. (2014). The purinergic neurotransmitter revisited: A single substance or multiple players?” Pharmacology & Therapeutics, 144(2), 162–191. doi:10.1016/j.pharmthera.2014.05.012.Find this resource:

Nässel, D. R., & Wegener, C. (2011). A comparative review of short and long neuropeptide F signaling in invertebrates: Any similarities to vertebrate neuropeptide Y signaling? Peptides, 32(6), 1335–1355. doi:10.1016/j.peptides.2011.03.013.Find this resource:

Nichols, D. E., & Nichols, C. D. (2008). Serotonin receptors. Chemical Reviews, 108(5), 1614–1641. doi:10.1021 /cr078224o.Find this resource:

Nichols, R. (2003). Signaling pathways and physiological functions of Drosophila melanogaster FMRFamide-related peptides. Annual Review of Entomology, 48 (January), 485–503. doi:10.1146/annurev.ento.48.091801.112525.Find this resource:

Nishimura, K., Kitamura, Y., Taniguchi, T., & Agata, K. (2010). Analysis of motor function modulated by cholinergic neurons in planarian Dugesia japonica. Neuroscience, 168(1), 18–30. doi:10.1016/j.neuroscience.2010.03.038.Find this resource:

Nishimura, K., Kitamura, Y., Umesono, Y., Takeuchi, K., Takata, K., Taniguchi, T., & Agata, K. (2008a). Identification of glutamic acid decarboxylase gene and distribution of GABAergic nervous system in the planarian Dugesia japonica. Neuroscience, 153(4), 1103–1114. doi:10.1016 /j.neuroscience.2008.03.026.Find this resource:

Nishimura, K., Kitamura, Y., Inoue, T., Umesono, Y., Yoshimoto, K., Taniguchi, T., & Agata, K. (2008b). Characterization of tyramine beta-hydroxylase in planarian Dugesia japonica: Cloning and expression. Neurochemistry International, 53(6–8), 184–192. doi:10.1016/j.neuint.2008.09.006.Find this resource:

Nishimura, K., Unemura, K., Tsushima, J., Yamauchi, Y., Otomo, J., Taniguchi, T., . . . Kitamura, Y. (2009). Identification of a novel planarian G-protein-coupled receptor that responds to serotonin in Xenopus laevis oocytes. Biological & Pharmaceutical Bulletin, 32(10), 1672–1677. this resource:

Niswender, C. M., & Conn, P. J. (2010). Metabotropic glutamate receptors: Physiology, pharmacology, and disease. Annual Review of Pharmacology and Toxicology, 50 (January), 295–322. doi:10.1146/annurev.pharmtox.011008.145533.Find this resource:

Noël, F., Mendonça-Silva, D. L., Thibaut, J-P B., & Lopes, D. V. S. (2007). Characterization of two classes of benzodiazepine binding sites in Schistosoma mansoni. Parasitology, 134(Pt 7), 1003–1012. doi:10.1017/S0031182007002442.Find this resource:

Novozhilova, E., Kimber, M. J., Qian, H., McVeigh, P., Robertson, A. P., Zamanian, M., . . . Day, T. A. (2010). FMRFamide-like peptides (FLPs) enhance voltage-gated calcium currents to elicit muscle contraction in the human parasite Schistosoma mansoni. PLoS Neglected Tropical Diseases, 4(8), e790. doi:10.1371/journal.pntd.0000790.Find this resource:

Ogasawara, M., Yamauchi, K., Satoh, Y.-I., Yamaji, R., Inui, K., Jonker, J. W., . . . Maeyama, K. (2006). Recent advances in molecular pharmacology of the histamine systems: Organic cation transporters as a histamine transporter and histamine metabolism. Journal of Pharmacological Sciences, 101(1), 24–30. this resource:

Omar, H. H., Humphries, J. E., Larsen, M. J., Kubiak, T. M., Geary, T. G., Maule, A. G., . . . Day, T. A. (2007). Identification of a platyhelminth neuropeptide receptor. International Journal for Parasitology, 37(7), 725–733. doi:10.1016/j.ijpara.2006.12.019.Find this resource:

Patocka, N., & Ribeiro, P. (2007). Characterization of a serotonin transporter in the parasitic flatworm, Schistosoma mansoni: Cloning, expression and functional analysis. Molecular and Biochemical Parasitology, 154(2), 125–133. doi:10.1016/j.molbiopara.2007.03.010.Find this resource:

(p. 243) Patocka, N., & Ribeiro, P. (2013). The functional role of a serotonin transporter in Schistosoma mansoni elucidated through immunolocalization and RNA interference (RNAi). Molecular and Biochemical Parasitology, 187(1), 32–42. doi:10.1016/j.molbiopara.2012.11.008.Find this resource:

Patocka, N., Sharma, N., Rashid, M., & Ribeiro, P. (2014). Serotonin signaling in Schistosoma mansoni: A serotonin-activated G protein-coupled receptor controls parasite movement. PLoS Pathogens, 10(1), e1003878. doi:10.1371 /journal.ppat.1003878.Find this resource:

Pax, R. A., Siefker, C., & Bennett, J. L. (1984). Schistosoma mansoni: Differences in acetylcholine, dopamine, and serotonin control of circular and longitudinal parasite muscles. Experimental Parasitology, 58(3), 314–324. this resource:

Peymen, K., Watteyne, J., Frooninckx, L., Schoofs, L., & Beets, I. (2014). The FMRFamide-like peptide family in nematodes. Frontiers in Endocrinologyi, 5 (January), 90. doi:10.3389 /fendo.2014.00090.Find this resource:

Pin, J.-P., & Acher, F. (2002). The metabotropic glutamate receptors: Structure, activation mechanism and pharmacology. Current Drug Targets. CNS and Neurological Disorders, 1(3), 297–317. this resource:

Purcell, J., Robertson, A. P., Thompson, D. P., & Martin, R. J. (2002). PF4, a FMRFamide-related peptide, gates low-conductance Cl(-) channels in Ascaris suum. European Journal of Pharmacology, 456(1–3), 11–17. this resource:

Putrenko, I., Zakikhani, M., & Dent, J. A. (2004). A family of acetylcholine-gated chloride channel subunits in Caenorhabditis elegans. Journal of Biological Chemistry, 280(8), 6392–6398. doi:10.1074/jbc.M412644200.Find this resource:

Ranganathan, R., Cannon, S. C., & Horvitz, H. R. (2000). MOD-1 is a serotonin-gated chloride channel that modulates locomotory behaviour in C. elegans. Nature, 408 (6811), 470–475. doi:10.1038/35044083.Find this resource:

Raouf, R., Blais, D., & Séguéla, P. (2005). High zinc sensitivity and pore formation in an invertebrate P2X receptor. Biochimica et Biophysica Acta, 1669(2), 135–141. doi:10.1016/j.bbamem.2005.01.009.Find this resource:

Rawls, S. M., Thomas, T., Adeola, M., Patil, T., Raymondi, N., Poles, A., . . . Raffa, R. B. (2009). Topiramate antagonizes NMDA- and AMPA-induced seizure-like activity in planarians. Pharmacology Biochemistry and Behavior, 93(4), 363–367. doi:10.1016/j.pbb.2009.05.005.Find this resource:

Reddien, P. W., Bermange, A. L., Murfitt, K. J., Jennings, J. R., & Alvarado, A. S. (2005). Identification of genes needed for regeneration, stem cell function, and tissue homeostasis by systematic gene perturbation in planaria. Developmental Cell, 8(5), 635–649. doi:10.1016/j.devcel.2005.02.014.Find this resource:

Ribeiro, P. (2015). Exploring the role of biogenic amines in schistosome host-parasite interactions. Trends in Parasitology, 31(9), 404–405. doi:10.1016/ this resource:

Ribeiro, P., El-Shehabi, F., & Patocka, N. (2005). Classical transmitters and their receptors in flatworms. Parasitology, 131 (Suppl. January), S19–S40. doi:10.1017 /S0031182005008565.Find this resource:

Ribeiro, P., & Patocka, N. (2013a). Neurotransmitter transporters in schistosomes: Structure, function and prospects for drug discovery. Parasitology International 62(6), 629–638. doi:10.1016/j.parint.2013.06.003.Find this resource:

Ribeiro, P., & Patocka, N. (2013b). Neurotransmitter transporters in schistosomes: Structure, function and prospects for drug discovery. Parasitology International 62(6): 629–638. doi:10.1016/j.parint.2013.06.003.Find this resource:

Ribeiro, P., & Webb, R. A. (1983). The synthesis of 5-hydroxytryptamine from tryptophan and 5-hydroxytryptophan in the cestode Hymenolepis diminuta. The International Journal for Parasitology, 13(1), 101–106.Find this resource:

Roeder, T. (2005). Tyramine and octopamine: Ruling behavior and metabolism. Annual Review of Entomology, 50(January), 447–477. doi:10.1146/annurev .ento.50.071803.130404.Find this resource:

Roeder, T., Seifert, M., Kähler, C., & Gewecke, M. (2003). Tyramine and octopamine: Antagonistic modulators of behavior and metabolism. Archives of Insect Biochemistry and Physiology, 54(1), 1–13. doi:10.1002/arch.10102.Find this resource:

Saberi, A., Jamal, A., Beets, I., Schoofs, L., Newmark, P. A. (2016). GPCRs direct germline development and somatic gonad function in planarians. PLoS Biology, 14(5), e1002457. doi:10.1371/journal.pbio.1002457.Find this resource:

Schoenfeld, B. P., Choi, R. J., Choi, C. H., Terlizzi, A. M., Hinchey, P., Kollaros, M., . . . McBride, S. M. (2013). The Drosophila DmGluRA is required for social interaction and memory. Frontiers in Pharmacology, 4(January), 64. doi:10.3389/fphar.2013.00064.Find this resource:

Scimone, M. L., Kravarik, K. M., Lapan, S. W., & Reddien, P. W. (2014). Neoblast specialization in regeneration of the planarian Schmidtea mediterranea. Stem Cell Reports, 3(2), 339–352. doi:10.1016/j.stemcr.2014.06.001.Find this resource:

Shimoyama, S., Inoue, T., & Kashima, M. (2016). Multiple neuropeptide-coding genes involved in planarian pharynx extension. Zoological Sciences, 33(3), 311–319.Find this resource:

Stark, H. (2007). Histamine receptors. Biotrend Reviews, 1(11), 1–8. this resource:

Sukhdeo, M. V., Hsu, S. C., Thompson, C. S., & Mettrick, D. F. (1984). Hymenolepis diminuta: Behavioral effects of 5-hydroxytryptamine, acetylcholine, histamine and somatostatin. The Journal of Parasitology, 70(5), 682–688. this resource:

Sukhdeo, S. C., Sangster, N. C., & Mettrick, D. F. (1986). Effects of cholinergic drugs on longitudinal muscle contractions of fasciola hepatica. The Journal of Parasitology, 72(6), 858–864. this resource:

Taman, A., & Ribeiro, P. (2009). Investigation of a dopamine receptor in Schistosoma mansoni: Functional studies and immunolocalization. Molecular and Biochemical Parasitology, 168(1), 24–33. doi:10.1016/j.molbiopara.2009.06.003.Find this resource:

Taman, A., & Ribeiro, P. (2011a). Characterization of a truncated metabotropic glutamate receptor in a primitive metazoan, the parasitic flatworm Schistosoma mansoni. PloS One, 6(11), e27119. doi:10.1371/journal.pone.0027119.Find this resource:

Taman, A., & Ribeiro, P. (2011b). Glutamate-mediated signaling in Schistosoma mansoni: A novel glutamate receptor is expressed in neurons and the female reproductive tract. Molecular and Biochemical Parasitology, 176(1), 42–50. doi:10.1016/j.molbiopara.2010.12.001.Find this resource:

Thompson, C. S., Sangster, N. C., & Mettrick, D. F. (1986). Cholinergic inhibition of muscle contraction in Hymenolepis diminuta (Cestoda). Canadian Journal of Zoology, 64, 2111–2115.Find this resource:

(p. 244) Tomosky, T. K., Bennett, J. L., & Bueding, E. (1974). Tryptaminergic and dopaminergic responses of Schistosoma mansoni. The Journal of Pharmacology and Experimental Therapeutics, 190(2), 260–271. this resource:

Tsai, I. J., Zarowiecki, M., Holroyd, N., Garciarrubio, A., Sanchez-Flores, A., Brooks, K. L., . . . Berriman, M. (2013). The genomes of four tapeworm species reveal adaptations to parasitism. Nature, 496(7443), 57–63. doi:10.1038 /nature12031.Find this resource:

van Nierop, P. (2005). Identification of molluscan nicotinic acetylcholine receptor (nAChR) subunits involved in formation of cation- and anion-selective nAChRs. Journal of Neuroscience, 25(46), 10617–10626. doi:10.1523 /JNEUROSCI.2015-05.2005.Find this resource:

Vogel, K. J., Brown, M. R., & Strand, M. R. (2013). Phylogenetic investigation of peptide hormone and growth factor receptors in five dipteran genomes. Frontiers in Endocrinology, 4(January), 193. doi:10.3389/fendo.2013.00193.Find this resource:

Vogt, K. (2015). Diversity in GABAergic signaling. Advances in Pharmacology, 73(January), 203–222. doi:10.1016 /bs.apha.2014.11.009.Find this resource:

von Kügelgen, I., & Hoffmann, K. (2015). Pharmacology and structure of P2Y receptors. Neuropharmacology, October. doi:10.1016/j.neuropharm.2015.10.030.Find this resource:

Wang, J., & Yu, Y. (2016). Insights into the channel gating of P2X receptors from structures, dynamics and small molecules. Acta Pharmacologica Sinica, 37(1), 44–55. doi:10.1038/aps.2015.127.Find this resource:

Williams, M. (2002). Purinergic neurotransmission. In K. L. Davis, D. Charney, J. T. Coyle, & C. Nemeroff (Eds.), Neuropsychopharmacology: The Fifth Generation of Progress (pp. 191–206), Philidelphia, Lippincott, Williams & Wilkins.Find this resource:

Wolstenholme, A. J. (2012). Glutamate-gated chloride channels. The Journal of Biological Chemistry, 287(48), 40232–40238. doi:10.1074/jbc.R112.406280.Find this resource:

Xu, G., Wu, S.-F., Wu, T.-S., Gu, G.-X., Fang, Q., & Ye, G. Y. (2015). De novo assembly and characterization of central nervous system transcriptome reveals neurotransmitter signaling systems in the rice striped stem borer, Chilo suppressalis. BMC Genomics, 16(1), 525. doi:10.1186 /s12864-015-1742-7.Find this resource:

Yamamoto, S., & Seto, E. S. (2014). Dopamine dynamics and signaling in Drosophila: An overview of genes, drugs and behavioral paradigms. Experimental Animals/Japanese Association for Laboratory Animal Science, 63(2), 107–119. this resource:

Zamanian, M., Agbedanu, P. N., Wheeler, N. J., McVeigh, P., Kimber, M. J., & Day, T. A. (2012). Novel RNAi-mediated approach to G protein-coupled receptor deorphanization: Proof of principle and characterization of a planarian 5-HT receptor. PLoS One, 7(7), e40787. doi:10.1371/journal.pone.0040787.Find this resource:

Zamanian, M., Kimber, M. J., McVeigh, P., Carlson, S. A., Maule, A. G., & Day, T. A. (2011). The repertoire of G protein-coupled receptors in the human parasite Schistosoma mansoni and the model organism Schmidtea mediterranea. BMC Genomics, 12(January), 596. doi:10.1186 /1471-2164-12-596.Find this resource:

Zatylny-Gaudin, C., & Favrel, P. (2014). Diversity of the RFamide peptide family in mollusks. Frontiers in Endocrinology, 5(January), 178. doi:10.3389/fendo.2014.00178.Find this resource:

Zimmermann, H., & Braun, N. (1999). Ecto-nucleotidases—Molecular structures, catalytic properties, and functional roles in the nervous system. Progress in Brain Research, 120(January), 371–385. this resource: