Show Summary Details

Page of

PRINTED FROM OXFORD HANDBOOKS ONLINE (www.oxfordhandbooks.com). © Oxford University Press, 2018. All Rights Reserved. Under the terms of the licence agreement, an individual user may print out a PDF of a single chapter of a title in Oxford Handbooks Online for personal use (for details see Privacy Policy and Legal Notice).

date: 22 October 2018

Neurotransmitters and Neuropeptides of Invertebrates

Abstract and Keywords

This chapter introduces working definitions of neuropeptides and neurotransmitters from the perspective of invertebrate physiological processes. Neuropeptides and neurotransmitters are intercellular chemical signaling agents used by all animals. Chemical signaling augments or substitutes for electrical communication in the nervous system. When these agents act as neurotransmitters, they convert electrical signals to chemical signals across the synapse. As hormones, they circulate from a site of release to act at a more distant site in the body of the organism. Neuropeptides and neurotransmitters are classified into these groups mostly on the basis of their molecular size. This article describes several neuropeptide superfamilies and their wide scope of actions in model invertebrates. The article also describes the main neurotransmitters used by invertebrates.

Keywords: invertebrate nervous system, neurotransmitter, neuropeptide, neuropeptide superfamilies, chemical signaling

Neurotransmitters and neuropeptides are two classes of molecules that animals with any form of a nervous systems use to send intercellular messages between parts of the nervous system and throughout the body. Chemical substances that communicate signals in the nervous system act either as hormones or neurotransmitters, or both. Hormones enter the circulation from a bona fide endocrine gland or other site to act on the nervous system at a distance from the place they were synthesized. Neurotransmitters fulfill roles in electrochemical communication by local release into the synaptic cleft from vesicles that contain the neurotransmitter in the presynaptic element of the synapse. Signaling molecules often act as both hormones and neurotransmitters, for example, vasopressin and serotonin.

Dale’s principle, formulated by Eccles (1957, 1964) in the late 1950s, stated the idea “one nerve utilises one transmitter” (Burnstock, 2014, p. 1). In the decades since, important discoveries have repeatedly refuted the idea embodied in Dale’s principle. It has been demonstrated that a single neurotransmitter, say L-Glutamate (L-Glu), can operate both in the central and peripheral nervous systems. It has been shown that more than one neurotransmitter and/or neuropeptide can be contained inside a synaptic vesicle for release into the synaptic cleft and act on the postsynaptic cell. The multiple signaling agents will act synergistically or antagonistically over morphological or temporal scales to increase the available repertoire of a nerve granted by its excitatory potential.

This chapter takes up the subject of invertebrate neurotransmitters and neuropeptides from a physiological perspective. It treats together a signaling agent’s actions as a hormone and as a neurotransmitter. This approach was adopted because there is less argument about designations as neurotransmitter or neuropeptide than there is about hormone versus neurotransmitter. Short-acting chemical agents that modulate intercellularly, rather than carrying the message synaptically between excitable cells, could be classified as hormones, neuromodulators, or paracrine agents. An example is ATP co-released with another neurotransmitter from synaptic vesicles that act on a glial membrane near the synapse (MacDermott et al., 1999). Since activation of tailored receptors in all these membranes often defines the effect observed, it is more important for our purposes to focus on the end effect.

Neuropeptides

Peptide hormones and neurotransmitters are believed to be the most ancient signaling molecules (Grimmelikhuijzen et al., 2002), and they are not limited to neuronal signaling, although neurons usually serve as either releasers or targets. Most active peptide neurotransmitters are 4–20 amino acids in length, derived from larger precursor hormones that are posttranslationally modified to form often multiple copies of a neuropeptide and more than a single active compound. This may provide redundancies, different potencies at receptors that bind them, or agents with different signaling emphases. Peptides circulate as neurohormones, are synaptically released as neurotransmitters, or both. Simultaneous functions as both hormone and neurotransmitter are the most common. Their diversity of actions allows multifunctionality of the same neural machinery, and it also attests to their longevity. Neuropeptides are broken down and their action terminated by peptidases at the extracellular membrane of the target.

Peptide receptors usually couple to G-proteins (Jekely, 2013). Known invertebrate peptide receptors activate the same second messenger machinery as in vertebrates (Sossin & Abrams, 2009). These authors point out that intracellular signal transduction pathways were in place very early in evolution. Nevertheless, identification of dedicated receptors in invertebrates as a whole lags somewhat behind their identification in vertebrates (Cardoso & Larhammar, 2014), in part because the physiological studies sometimes have not been done. Neuropeptide signaling controls metabolic states such as somatic growth and condition as well as reproductive status, water, and salt balance. Peptide signaling molecules also mediate integration of the sensory environment into the homeostatic conditions of the animal, as in the establishment of clock rhythms.

The role of peptides in mediating invertebrate physiological processes has been studied since the early part of the last century, with the most complete and detailed information emerging from arthropod insect models. Studying the physiological actions of invertebrate peptides in noninsect models has been spotty and largely limited to crustaceans and mollusks. Bioinformatics processing of high throughput sequencing data is beginning to identify peptides plus their receptors in invertebrate phyla based on percent identity to known arthropod proteins. The establishment of physiological roles beyond insects is beginning in these phyla, and these efforts will more fully describe their actions throughout the Metazoa.

The annual special paper collection Invertebrate Neuropeptides (I through XV) published in the journal Peptides since 2001 and edited by Ronald J. Nachman as an outcome of the annual International Invertebrate Neuropeptide Conference is the seminal source of recent collected research on this topic, and the reader is referred there (http://www.journals.elsevier.com/peptides/special-issues) for the latest as well as the classical on the reader’s invertebrate species of choice.

Grouping neuropeptides into families based on structure-function is both a rational impulse and critical to understanding their effects at defined or putative receptors. Considerable super- and subcategorical complexity arises due to prohormone processing into numerous related peptides with sometimes contrasting function. In a pattern found repeatedly in the neuropeptides families, some members have inhibitory effects on metabolic pathways that other members stimulate. This often involves pleiotropy, in which the products of a single gene have unrelated phenotypic characteristics, although superfamilies commonly include numerous genes. Furthermore, the number of repeats change or the sequences diverge into distinct peptides over time (Webster, et al. 2012). Variable naming styles across the invertebrates further complicates the cataloging chore. It is pertinent that peptide names sometimes imply a narrow functional role. Although their actions can be specific, peptides are often named for the first physiological effect noted. Once, and if, wider functions are discovered for the peptide, the name, now an artifact of the originally studied function, can be misleading. These names persist, perhaps because the iconic physiology that defined them was executed so long ago (>100 years in some cases). Another complication is the designation as peptide hormone versus neurotransmitter. The scope of influence of these messengers across the brain and body is not sufficiently straightforward that this is always a clear distinction for a specific peptide. Thus, it can be up to the author to decide, as I have done occasionally here.

Jékely (2013) and Bauknecht and Jékely (2015) recently addressed some gaps and confusion among the categories of invertebrate neuropeptides and their receptors. Their work bridged the organizational distance between better-studied insects and other invertebrates, and due to the emphasis on noninsect invertebrate models here, it served as the categorical treatment of neuropeptides presented. Table S6 from their 2015 work is reprinted here as Figure 1.

Neurotransmitters and Neuropeptides of InvertebratesClick to view larger

Figure 1 The distribution by phylum of superfamilies of invertebrate neuropeptides, reprinted from Bauknecht and Jekely (2015). The red-underlined groups are discussed in the section on neuropeptides.

(This figure is a derivative of “Large-scale combinatorial deorphanization of Platynereis neuropeptide GPCRs,” www.cell.com/cell-reports/pdfExtended/S2211-1247(15)00678-6, which has been modified from Figure S6 of that work under the Creative Commons Creative Commons Attribution License [CC BY].)

Even with categorization, neuropeptides have diverse impacts on biological systems and occasionally appear highly tailored to species, even when their physiological role as a member of a peptide family is conserved. An example is a phosphorylated adipokinetic hormone (AKH) in the beetle Trichostetha fascicularis, whose role is to regulate carbohydrate supply during flight, in this species alone (for now; Gäde et al., 2006). The better we are at identifying neuropeptides and the genes from which they originate, however, the smaller the set of species-specific peptides becomes. Nevertheless, comprehensive summaries of their actions are a daunting task. Neuropeptide complexity and impact are illustrated, however, from consideration of several examples. The first two are examples of central nervous system peptide hormone command function in invertebrates-only neuropeptide groups, the crustacean hyperglycemic hormone (CHH) superfamily and the FMRFamides, while the others are important peptide families across Metazoa.

CHH Superfamily

The arthropod eyestalk X organ-sinus gland is part of the protocerebrum and a source of potent peptide hormones of the crustacean hyperglycemic hormone superfamily (CHH; eighth group from the top on the right side of Fig. 1). This neurohumeral analog of the vertebrate hypothalamus-pituitary system illustrates how centrally released peptides execute command functions, while related compounds as differently spliced proteins or from similar genes have narrower roles. The 80 or so neuropeptide products of approximately 25 CHH genes released from the eyestalk X organ-sinus gland are subgrouped into CHH type I, “true” CHH and ion transport proteins (ITP; Christie et al., 2010) and CHH type II, which are molt-inhibiting (MIH) and gonad-inhibiting hormones (GIH also referred to vitellogenin-inhibiting VIH), but also includes mandibular organ-inhibiting hormone (MOIH), which inhibits secretion of methyl farnesoate from the mandibular organs (Cary et al., 2011; Webster et al., 2012). The CHH peptides are uncommonly large, with type I neuropeptides consisting of 72 amino acids, while type II are even larger at, usually, 78 amino acids. All have structural features in common, especially the three disulphide bonds formed by the characteristic position of six cysteines.

The CHH peptides’ signature role is to regulate carbohydrate metabolism, that is, glycogen mobilization (Christie et al., 2010; Webster, 2012), responding rapidly at times of stress such as during toxicant exposure. Paradoxically, none of the known actions of type II CHH are hyperglycaemic. The effects of the first identified CHH member were of MIH, discovered >100 years ago when Zeleny (1905) noted that eyestalk removal precipitated molting.

An example of how the CHH family resources are marshaled in the bodies of arthropods is provided by insect and crustacean life histories surrounding the periodic molt, or ecdysis. MIH from the X organ-sinus gland inhibits transition to preecdysis by preempting molting hormone (MH) release from the eyestalk Y organ. Release of MH precipitates molting, and once MH is released, the Y organ is refractory to MIH (Chung & Webster, 2003; Nakatsuji & Sonobe, 2004; Nakatsuji et al., 2006). Thus, MIH has master control of elements of the intermolt period up to premolt (Passano, 1953). Critically, CHH is molt-inhibiting in penaeids, lobsters, and crayfish (Chang et al., 1990), but it appears that in crustaceans that preferentially use MIH for this role, CHH also inhibits the molt, with much lower potency (Wainwright et al., 1996).

The main function of CHH in exercise or stress is to mobilize glycogen by increasing glycogen synthase activity, resulting in hyperglycemia in hepatopancreas and muscles. CHH has other nutrient-marshaling actions in that it also stimulates the release of amylase from the midgut gland (Sedlmeier, 1988) and raises titers of phospholipids and free fatty acids (Santos et al., 1997). CHH release and hyperglycemia normally is episodic on a time scale of minutes, with glucose inhibiting the release of CHH in a negative feedback process (Santos & Keller, 1993; Glowik et al., 1997). This energetic function for CHH is not specifically tied to either intermolt or molting, although the molt is a life-threatening stress in the life of arthropods that calls upon glycogen stores (Gäde, 2009). CHH released from the gut, however, causes water absorption to the gut critical to ecdysis (Chung et al., 1999); this is also the role of the CHH family member ITP in insects of xeric environments (Audsley et al., 1992). Last, in pleiotropy of water absorption, a CHH isoform can increase the osmotic concentration of hemolymph, for example, during hyposalinity exposures (Dircksen et al., 2001).

MIOH inhibits the release of methyl farnesoate from the mandibular organs. Methyl farnesoate is a form of juvenile hormone in both insects and some crustaceans (Wainwright et al., 1996; Nagaraju, 2007). Thus, its inhibition precipitates reproductive maturity, for example, oocyte maturation (Jo et al., 1999).

FMRFamide Family

The FMRFamide family of neuropeptides (top left family in Fig. 1) containing Arg-Phe-NH2 at the carboxyl terminus is the product of a single gene (Nässel, 1996). The FMRFamides’ generalized role in invertebrates is control of synaptic transmission at neuromuscular junctions, and muscle contractions (Milakovic et al., 2014). The different mRNA transcripts produced from the FMRFamide gene contain sequences for tetra-, hepta-, and hexapeptides. Expression of one of the different family members is often prioritized in different neurons, and innervation by multiple neurons exerts pleiotropic effects on the same muscle (Van Golen et al., 1995). In other neurons more than one of these peptides occurs. The FMRFamide neuropeptides are used centrally and in motor neurons that control heartbeat, respiration, egg laying, and copulation (Santama et al., 1995; Van Golen et al., 1995).

GnRH Superfamily

The GnRH superfamily (tenth family from top left of Fig. 1) is named for the approximately 30-member gonadotropin-releasing hormones of vertebrates (GnRH-I, and –II, also –III in teleosts; Kah et al., 2007; Tsai & Zhang, 2008), whose activation of the hypothalamo-pituitary-gonadal axis initiates reproductive behavior and supresses feeding (Temple et al., 2003; Kauffman & Rissman, 2004; Matsuda et al., 2008).

GnRH induces gonadal steriodogenesis and gametogenesis in vertebrates via its stimulation of secretion of luteinizing hormone (LH) and follicle-stimulating hormone (FSH). GnRH is also prominent in invertebrates, where it has many actions supporting reproduction, as well as other effects. GnRH is 10–12 amino acids in all animals (Guan et al., 2014). Invertebrate GnRH family members have two additional N-terminal residues compared to vertebrates, but they lack others at the C-terminus (Tsai, 2006). Useful amino acid alignments of both vertebrate and invertebrate GnRH were given by Lindemans et al. (2011).

The GnRH family also is strongly represented in invertebrates by the AKH known from insects, cladocerans such as Daphnia, marine crustaceans like H. americanus and Cancer borealis, and the opisthobranch A. californica, amid perhaps other mollusks for which sequence information was collected (Iwakoshi-Ukena et al., 2004; Roch et al., 2011; Johnson et al., 2014). Red pigment-concentrating hormones (RPCH) and corazonin (CRZ) round out the invertebrate representatives in the GnRH superfamily.

GnRH has broad and pleiotropic actions in invertebrates in promoting/inhibiting reproduction and in contracting/relaxing skeletal and smooth muscle. GnRH is expressed widely throughout invertebrate nervous systems. Octopus vulgaris GnRH (oct-GnRH) increased steroidogenesis in gonads and induced oviduct contraction in addition to stimulating cardiac output (Iwakoshi-Ukena et al., 2004; Kanda et al., 2006). In A. californica, Ap-GnRH is expressed most highly in pedal ganglia, whose motor neurons control the muscles of the parapodia. GnRH caused parapodial relaxation (Johnson et al., 2014). Ap-GnRH also inhibited feeding and promoted attachment to the substrate (Tsai et al., 2010; Johnson et al., 2014); both behaviors precede egg-laying (Macginitie, 1934; Strumwasser et al., 1969; Pennings, 1991). Ap-GnRH inhibited and reversed gonadal maturation and promoted elimination by promoting intestinal contractions. Both actions were shared with AKH in this species (Johnson et al., 2014). In Ciona intestinalis GnRH modulated the gonadal release and synthesis of testosterone and progesterone (D’Aniello et al., 2003).

AKH and RPCH of invertebrates have similar structure. AKH synthesis, release, binding, and actions are fully described (Gäde, 2009; Johnson et al., 2014). The decapeptide AKH is important to carbohydrate and lipid energy mobilization in insects (Bednářová et al., 2013), especially such as during flight (Gäde, 2009). AKH released from the corpora cardiaca gland in the insect brain activates a stimulatory G(q) protein linked to a triacylglycerol lipase or a glycogen phosphorylase in the fat body, where carbohydrates are stored. The lipids are broken down to monoacylglycerols and released into the hemolymph. RPCH control and modulate gut motility via effects in the stomatogastric ganglion (Nusbaum & Marder, 1988; Dickinson et al., 1993, 1997; Johnson et al., 2014), as well as controlling red pigment in the eye (see later; Ranga Rao & Riehm, 1988). CRZ, named for its original purpose in elevating heart rate (Veenstra, 1989), may have actions related to energy budgeting in response to inadequate nutrition (Veenstra, 2009); it has been demonstrated to control insect body color, and it influences the proportional growth of different body segments (Sugahara et al., 2016; Tanaka et al., 2016).

The photonic energy reaching the rhabdomes of each ommatidium of compound eyes of arthropods can be controlled by the fore and aft movement of pigment granules that allow various amounts of light into the ommatidia and control the spectral sensitivity (Autrum, 1981; Ranga Rao & Riehm, 1988; Meyer-Rochow, 2001; Meelkop et al., 2011). The light-adapted scenario has pigments dispersed and thus kept from reaching the rhabdomes, or they can be aggregated in dark-adapted eyes. Pigment movement is controlled by pigment-dispersing and pigment-concentrating hormones, among which are RPCH. The pigment movement hormones, in turn, are released by neurotransmitters. For example, in Uca pugilator, norepinephrine (NE) elicited a light-adapting response, while dopamine (DA) caused a dark-adapting response (Kulkarni & Fingerman, 1986).

Vasopressin Superfamily

Vasopressin and oxytocin (VP and OT; eighth family from top left of Fig. 1) belong to a large superfamily 650 million years old, with strong evolutionarily conservation (Donaldson & Young, 2008; Goodson, 2008). A common ancestral gene in vertebrates underwent duplication in a jawed ancestor (Goodson, 2008) so that these peptides are the products of different genes. Invertebrates, with cephalopods as an exception, have a single gene family homolog (Donaldson & Young, 2008).

The VP superfamily plays two important physiological roles in vertebrates, regulating the hypothalamic-pituitary-adrenal axis, especially related to osmoregulatory and diuretic control at the kidneys, and protecting cerebral blood flow (McEwen, 2004). VP and OT regulate vascular blood pressure via controlling the release of nitric oxide (Katusić, 1992), among other actions, dilating some vessels and constricting others. VP and OT are also social hormones in vertebrates, promoting maternal closeness, are important in social recognition, and mitigate stress (Donaldson & Young, 2008; Wójciak et al., 2012). The structure is a nonapeptide, a covalent ring structure closed by a disulfide bridge, with a flexible tail of three residues, with VT and OT differing by just two residues.

The VP superfamily has been understudied in invertebrates. The osmoregulatory and fluid balance role of VP is conserved in phyla from nematodes to tunicates (van Kesteren et al., 1995; Satake et al., 1999; Kawada et al., 2004; Ukena et al., 2008; Minakata, 2010; Sakamoto et al., 2015). Gruber (2014) summarized functional studies in insects, annelids, gastropods, cephalopods, and C. elegans. Of special note among the roles of VP superfamily members apart from the diuretic and antidiuretic effects is a role in learning in Sepia officinalis and C. elegans (Bardou et al., 2010; Beets et al., 2012; Garrison et al., 2012); a role in reflex behaviors in A. californica (Martínez-Padrón et al., 1992); and a reproductive role in earthworm and the gastropod Lymnaea stagnalis (van Kesteren et al., 1995; Ukena et al., 1995; Oumi et al., 1996; Fujino et al., 1999).

Other important neuropeptide families in invertebrates that have vertebrate counterparts are the growth inhibitory and cardiac activity-modulating somatostatin/allostatins (AST), the allatotropins (AT), which largely oppose the effects of AST on growth, and gastrin/cholecystokinin (CCK)/sulfakinin (SK), whose main roles are associated with inhibition of feeding.

Neurotransmitters

To be defined as a neurotransmitter, a chemical substance must fulfill all of the following critical criteria. It must be present within the synapse, with the machinery for both its synthesis and breakdown also present there, and it must bind and have an action in the synapse (Purves et al., 2001). Refinements of these categories have arisen to require that the substance be released by nerve stimulation, its application to the synapse must mimic the postsynaptic effects of presynaptic stimulation, and its actions must be prevented by an established receptor blocker. By these criteria, L-Glutamate (L-Glu), the major excitatory neurotransmitter in the brains of all metazoans, could not be defined as a neurotransmitter at ionotropic glutamate receptors (iGluR) until the early 1980s (Lee Johnson, 1978; Davies & Watkins 1982a,b; Watkins 1981a,b; Watkins & Evans, 1981; Mayer & Westbrook, 1987). This failure was based primarily on the paucity of specific blockers of the iGluR that could help identify the iGluR subtypes whose actions upon agonist binding were often not uniform. Without the ability to block specific actions of L-Glu at what became known as the subfamilies of N-methyl-D-aspartate (NMDA) and non-NMDA (AMPA- and quisqualate-activated, and kainite) receptors, it could not be unequivocally demonstrated that effects were induced by L-Glu.

One basis for separating neurotransmitters from neuropeptides in classifying communication styles in the nervous system is by size. Neurotransmitters are smaller than neuropeptides, and they are sometimes single amino acids, such as L-Glu, NMDA, and gamma amino butyric acid (GABA). Other small molecule neurotransmitters are NE (also called noradrenaline), epinephrine (EPI), serotonin (5-HT), histamine (HA), and acetylcholine (ACh). One rule of thumb is that if the signaling molecule is larger than three amino acids, it is a neuropeptide (Purves et al., 2001).

We have already described some neuropeptide families that are either demonstrated or putative neurotransmitters used by invertebrates. Invertebrate phyla also use nonpeptide neurotransmitters at receptors that high throughput sequencing techniques are demonstrating are often quite similar to those used by vertebrates. Important in invertebrates are amino acid neurotransmitters L- and D-Glu, D- and L-aspartate (Asp), GABA, glycine, and D-serine. Amine neurotransmitters of invertebrates are HA; the catecholamines DA, NE, and EPI; 5-HT; melatonin; octopamine (OA); and tyramine. A gaseous neurotransmitter is nitric acid (NO). ACh is a small, unique molecule that has perhaps the longest reputation as a defined neurotransmitter, thanks to the frog neuromuscular junction. Most invertebrate model species have all of these as well as a full complement of neuropeptides.

There is one enormous exception, and that is the Phylum Ctenophora. Ctenophores have been proposed as the sister group to all other metazoan animals, based on profound differences in their nervous system for which there is no role for most small molecule neurotransmitters, except L-Glu acting at as many as eight candidate iGluR (Ryan et al., 2013; Moroz & Kohn, 2015, 2016). Aside from L-Glu, none of the critical criteria for traditional neurotransmitters can be filled for ctenophores, although this presumption is based largely on consideration of the genome, in addition to some targeted physiological studies (Moroz & Kohn, 2015), rather than on the basis of extensive physiological studies using all conceivable agonists. Ctenophores express >100 peptide-receptor-like G proteins plus other peptide-receptor like proteins, however, suggesting that neuropeptide signaling may be quite diverse and may convey fast excitatory neurotransmission in the ctenophores (Moroz et al., 2014).

The slightly different architecture of the invertebrate synapse from vertebrates can make it tricky to study the effects of neurotransmitters. When the synapse occurs in the neuropil connecting ganglia, and not near the cell body, the synapse cannot easily be studied by isolating the pre- or postsynaptic neuronal cells (Brown & Piscopo, 2013). When it is not possible to study a synapse electrophysiologically, the chemical presence of either a neurotransmitter or part of its machinery for synthesis or breakdown can be used to indicate the neurochemical nature of the synapse. Thus, although famous invertebrate models tend to be the species in which detailed information is available on the physiological actions of neurotransmitters, stereotyping of neurotransmitter actions can be extrapolated to understudied invertebrate phyla based on these chemical signatures.

ACh

ACh mediates fast excitatory cholinergic neurotransmission in the nervous system of all metazoans that use this neurotransmitter. These actions are conveyed via ACh-induced activation of pentameric nicotinic receptors, which are archtypical members of the ligand-activated receptor superfamily that includes 5-HT-, GABA-, and glycine receptors (Sattelle et al., 2005). ACh is synthesized in nerve terminals from acetyl coenzyme A and choline, by the enzyme choline acetyltransferase (CAT), and broken down by acetylcholine esterase (AChE) after reuptake into neurons and glial cells (Banks et al., 2009) by transporters.

Neuromuscular transmission in invertebrates is both glutamatergic (Hooper et al., 1986; Stein et al., 2006) and cholinergic (Futamachi, 1972; Marder, 1974, 1976; Weiss et al., 1992; Brezina et al., 1995; Katz & Frost, 1996; Kratsios et al., 2012), a trait shared with vertebrates (Vyas & Bradford, 1987; Brunelli et al., 2005; Pizzi et al., 2006; Rinholm et al., 2007). Visceral motor activity in crustaceans is commanded by the stomatogastric ganglion and thus is largely cholinergic (Gallus et al., 2006). Independent confirmation of the cholinergic control of gut motility has been found in mollusks and annelids, although numerous other neurotransmitters and neuropeptides participate (Anctil et al., 1984; Lloyd & Willows, 1988; Ukena et al., 1995). In nematodes, cholinergic neurotransmission controls the chemosensory nerve output of the amphids, a pair of cephalic sensory organs, and pharangeal pumping is under cholinergic control (Croll, 1977).

Much of cholinergic neurotransmission is inhibitory, increasing Cl or K+ conductances in postsynaptic neurons. This has been studied in detail in Aplysia (Kehoe, 1972a,b; Inoue et al., 1994; Kehoe & Vulfius, 2000; and many others).

ACh also activates muscarinic receptors associated with G proteins, although these are not well studied in invertebrates (Liu et al., 2016); most studies are in insects. Muscarinic receptors located presynaptically were found to inhibit release of ACh and other neurotransmitters in a form of negative feedback, whereas those located on the postsynaptic membrane induced excitatory responses both centrally and peripherally (Trimmer, 1995; Caulfield & Birdsall, 1998).

L-Glu

L-Glu is the fast excitatory neurotransmitter of the central nervous system of all Metazoa. This non-essential dietary amino acid is synthesized locally in neurons from glutamine by glutaminase. Another source is 2-oxoglutarate, an intermediate of the tricarboxylic acid cycle. L-Glu is taken up from the synaptic cleft by transporters in presynaptic terminals and glia. In vertebrate glia, L-Glu is converted into glutamine by glutamine synthetase for transport back to the presynaptic neuron, and there is scant evidence that this occurs in crustaceans (Sullivan et al., 2007).

As mentioned above, L-Glu is a critical neurotransmitter at the neuromuscular junction in all invertebrates. There is growing evidence that the iGluR subfamilies so important in vertebrates are also present in invertebrates, although the protein sequence similarity of putative NMDA-, AMPA- and kainite-like receptors to those of vertebrates is not sufficient for specific receptor effects to be isolated or defined with much precision. Nevertheless, L-Glu and its congeners such as NMDA and D-Asp have as widespread actions as ACh in excitable neurotransmission in invertebrates. Glutamatergic agonists cause contraction of striated muscle (Usherwood et al., 1984), and convey sensory information to central ganglia (Bravarenko et al., 2003; Kempsell & Fieber, 2015) among many other specific physiological actions.

L-Glu also can have inhibitory actions such as activation of Cl– conductances in pentameric ligand-activated receptors in nematodes and insects (Fuse et al., 2015).

Amines

The catecholamines DA, NE, and EPI all derive from tyrosine (Purves et al., 2001), as do OA and tyramine. As neurotransmitters, they are taken up from the synaptic cleft by Na+-dependent transporters. 5-HT is made from tryptophan. HA is made from histidine. Invertebrate amine neurotransmitters are primarily DA, OA, tyramine, HA, and 5-HT.

OA and tyramine are probably unique to invertebrates as neurotransmitters and hormones. OA acts analogously to vertebrate epinephrine and norepinephrine (Verlinden et al., 2010); all four are derived from tyrosine and thus chemically closely related. OA, at least, is enormously important with diverse actions. Tyramine is much less studied. Each has been most comprehensively studied in insects. They appear to activate G-protein-coupled receptors associated with stimulation or inhibition of adenylyl cyclase activity, Gs and Gi, respectively (Blenau & Baumann, 2001).

OA induces release of AKH from the central nervous system as well as induces release of fatty acids from the fat body, and thus it has a critical role in energy mobilization (Verlinden et al., 2010). It initiates the stress response as the fight-or-flight hormone of invertebrates (Blenau & Baumann, 2001). OA controls the activity of flight muscles. It increases the sensitivity of sensory receptors in the periphery through a complex of effects such as modifications of neuronal membrane resistance, adaptation of receptors to stimuli, enhancement of postsynaptic responsiveness, and the shape of the action potential. It also triggers ovulation and plays a critical role in olfactory conditioning in the honeybee.

In C. elegans, tyramine had a role in pathogenic avoidance learning (Jin et al., 2016). It has been suggested that tyramine may oppose the effects of OA by inhibiting adenylyl cyclase through a Gi-like receptor.

In the crustacean eye, NE, acting at α-adrenoreceptors, light-adapts distal retinal pigment by stimulating the release of light-adapting hormone, actions directly opposed by DA and its induction of dark-adapting hormone release (Fingerman et al., 1994). HA shepherds partially dark-adapted distal retinal pigments to their fully dark-adapted state by inhibiting the release of light-adapting hormone, while preventing partially light-adapted pigments from moving to fully light adapted (Stuart et al., 2007).

5-HT and OA, acting hormonally in release from the pericardial organ, a neurohemal structure, increase the crustacean heartbeat by modulating output of the cardiac ganglion; they also increase cardiac myocyte contractility. DA and NE act similarly in the heart, increasing heartbeat rate and amplitude via effects on cardiac ganglion neuronal bursting activity, stimulating or inhibiting different neurons. DA acts at Gs- and Gi-coupled receptors in this capacity, as well as in other actions in the NS (Blenau et al., 1998; Barbas et al., 2006). DA also affects heartbeat by modulating output of the interneurons the form the cardiac pacemaker. HA suppresses cardiac ganglion motor neuron activity by activating a chloride conductance.

The crustacean stomatogastric ganglion innervates the muscles of the foregut and is connected to the rest of the central nervous system via the stomatogastric nerve; its two groups of neurons control gastric mill muscular contractions and contractions of the pyloric stomach muscles, respectively. DA and to a lesser extent 5-HT and OA enhance nerve-evoked contractions of the foregut muscle. 5-HT modulates the pyloric muscle contraction rhythm by either postsynaptic or hormonal effects on the muscle. In Aplysia, DA- and 5-HT-mediated modulation of the buccal ganglia mediates ingestion and swallowing, respectively (Kabotyanski et al., 2000). Barbas et al. (2006) showed that 5-HT is a diversely acting molecule in invertebrates for which there are approximately five receptors in Aplysia, four in Drosophila, and three in C. elegans, with at least nine that can be classified with vertebrate 5-HT receptors; the rest have low similarity (Tierney, 2001). Serotonergic neurosecretory neurons dispense 5-HT into the lobster circulation to instigate agonistic behaviors, which are opposed by OA (Kravitz, 2000). Circulating 5-HT controls swimming behavior in the medicinal leech, Hirudo medicinalis, by activating the swim gating neurons (Kristan et al., 2005). 5-HT plays a large role in neural development in Drosophila and other insects (Blenau & Thamm, 2011).

5-HT induces dispersion of red, black, and/or white pigment in chromatophores in the crustacean cuticle; it acts as a neurotransmitter at 5-HT receptors to release red or black pigment movement hormones (RPDH and BPDH; Fingerman et al., 1994). DA acts either in mimic of these effects, or it inhibits them in some species, indirectly, like 5-HT, by controlling the release of pigment movement hormones. NE acts directly on melanophores via a-adrenoreceptors to disperse black pigment in the cuticle, while its effects on dispersing red pigment are via stimulation of release of RPDH. HA acting at H2 receptors inhibits BPDH release. OA also blocks release of BPDH.

5-HT and its precursor 5-hydroxytryptophan stimulate the release of MIH from the eyestalk X organ, culminating in suppression of ecdystone secretion from the Y organ (Fingerman et al., 1994). 5-HT and OA both partially block the release of methyl farnesoate from mandibular organs. 5-HT can stimulate ovarian development by triggering the release of gonad-stimulating hormone.

5-HT and OA acting hormonally activate a central motor program for postural flexion and postural extension in decapods, respectively, presumably by contrasting effects on the excitatory postsynaptic potential at the synapse of premotor interneurons with postural motor neurons (Fingerman et al., 1994). The amine neurotransmitters have numerous neuromuscular transmission effects in walking and swimming legs, and the tails and other appendages of decapods, but a uniting theme is that the receptors on motor neurons are usually for 5-HT, DA, NE, and OA.

In osmoregulatory control at the gills, DA and OA have contrasting effects on cAMP levels and thus effects on the Na-K-ATPase that acclimate euryhaline crustaceans to seawater (Fingerman et al., 1994). DA and 5-HT appear to control this process in acclimation to hypoosmotic environments.

5-HT mimics the effect of CHH in elevating hemolymph glucose, as do OA, NE, and EPI. Olfactory cells are rich in H2 receptors whose activation by HA suppresses activity. Some mechanoreceptors and stretch receptors in the appendages are sensitive to 5-HT and/or OA.

Finally, 5-HT plays strong roles in learning in Aplysia, mediating short-, intermediate-, and long-term memory of withdrawal reflexes (Bailey & Kandel, 2008; Lee et al., 2008; see also Chapter by Rankin on Nonassociative Learning in Invertebrates). Short-term memory induced by brief exposure to 5-HT is due to the modulation of a number of membrane currents (Byrne & Hawkins, 2015), one of which is a K+ current, whose reduction by PKA-dependent phosphorylation leads to depolarization and an increase in membrane excitability (Siegelbaum et al., 1982). 5-HT also activates protein kinase C, mobilizing transmitter and resulting in more release of neurotransmitter (L-Glu) into the synaptic cleft (Byrne & Hawkins, 2015). The overall effect is heightened excitability and increased neurotransmission using existing equipment in the synapse. Longer-term memory, induced by greater temporal exposure to 5-HT, on the other hand, involves new protein synthesis.

GABA and Glycine

GABA and glycine have reputations in the nervous system of both vertebrates and invertebrates as inhibitory neurotransmitters that activate ionic currents for chloride. GABA- and glycine-induced presynaptic inhibition are widespread phenomena in invertebrates (Gingl et al., 2004; Nishino et al., 2010). The axons of mechanosensory neurons in the periphery of the spider Cupiennius salei are especially vulnerable to GABA-induced inhibition, while in crustaceans stretch receptor neurons have GABAergic inhibition (Elekes & Florey, 1987). GABA can be excitatory at sensory neuron afferents in which the [Cl] is high. Glycine can be a coagonist at excitatory NMDA receptors, although its high concentration in seawater makes this less likely in marine invertebrates (Carlson et al., 2012).

ATP

The purines ATP and adenosine, plus pyrimidines, are believed ancient signaling molecules coopted very early by the nervous system from their roles in energy production (Burnstock & Verkhratsky, 2009). ATP and its metabolites are present in presynaptic vesicles due to the necessity for active transport to concentrate neurotransmitter into the vesicle. Thus, ATP and adenosine are coagonists with other ligands, activating nonspecific cation channels, or Ca2+ channels, to induce modulatory, often excitatory effects. Their receptors often activate adenylyl cyclase-linked G proteins. Some of the wide-reaching effects of purinergic receptor activation are modulation of muscle (mollusks) and cilia movement (ctenophores, embryonic echinoderms), including modulation of cardiac contractility (mollusks, arthropods), repair of mechanoreceptors (cnidarians), olfaction and gustation (arthropods), axon regeneration (annelids), inhibition of sexual maturation (echinoderms), and control of luminescence (echinoderms). Adenosine often elicits an inhibitory effect when it binds to specific presynaptic receptors.

NO

NO-mediated events are important in learning in gastropods (Kemenes et al., 2002; Susswein & Chiel, 2012; Korshunova & Balaban, 2014) and in long-term plasticity in Octopus ventral lobe (Shomrat et al., 2015). NO-induced plasticity may have been selected for in invertebrates (Moroz & Kohn, 2011) rather than, or in addition to, a system based on NMDA receptors for L-Glu, as in mammals (Shomrat et al., 2015).

Concluding Remarks

We have been introduced to the idea that the Metazoan nervous system may have evolved multiple times, with species with disparate life histories and physiologies differently emphasizing the ways in which signals are communicated from neural command centers to the periphery and back again. We considered here just a few of the more than 70 neuropeptide families and the 10 or so broad classifications of smaller-molecule neurotransmitters available. This diversity of raw materials, combined with their broad spectrum of effects in different species, makes evident that ample methods exist for conducting the work of a nervous system in invertebrate animals. As Webster (2012) noted, the pleiotropic and overlapping effects of neuropeptides and neurotransmitters within a family constitute challenges to deducing the principal biological function of these signaling molecules in a species, let alone in a class or phylum. There is so much yet left to discover, with sensitive molecular and phylogenetic studies now outpacing the important confirmatory studies of receptor binding and physiological actions. The evolutionary focus of the genetic studies, fortuitously, however, has and will continue to allow reinterpretation of the findings of older studies in this newly supported context. The evolutionary focus is also an exciting perspective for new studies on established model animals. Invertebrate models of nervous system physiology will always offer insights on how the complexity of vertebrate nervous systems came into existence. With or without a sophisticated central nervous system, neuropeptides and neurotransmitters are the constants and the foundation of any number of evolved nervous system designs.

References

Anctil, M., Laberge, M., & Martin, N. (1984). Neuromuscular pharmacology of the anterior intestine of Chaetopterus variopedatus, a filter-feeding polychaete. Comparative Biochemistry and Physiology C, 79, 343–351.Find this resource:

Audsley, N., McIntosh, C., & Phillips, J. E. (1992). Isolation of a neuropeptide from locust corpus cardiacum which influences ileal transport. Journal of Experimental Biology, 173, 261–274.Find this resource:

Autrum, H. (1981). Light and dark adaptation in invertebrates. In H. Autrum (Ed.), Handbook of sensory physiology, vol. VII/6C, Comparative physiology and evolution of vision in invertebrates (pp. 1–91). Berlin, Germany: Springer.Find this resource:

Bailey, C. H., & Kandel, E. R. (2008). Synaptic remodeling, synaptic growth and the storage of long-term memory in Aplysia. Progress in Brain Research, 169, 179–198.Find this resource:

Banks, G., Kemenes, I., Schofield, M., O’Shea, M., & Korneev, S. A. (2009). Acetylcholine binding protein of mollusks is unlikely to act as a regulator of cholinergic neurotransmission at neurite-neurite synaptic sites in vivo. FASEB Journal, 23, 3030–3036.Find this resource:

Barbas, D., Zappulla, J. P., Angers, S., Bouvier, M., Mohamed, H. A., Byrne, J. H., … DesGroseillers, L. (2006). An Aplysia dopamine1-like receptor: molecular and functional characterization. Journal of Neurochemistry, 96, 414–427.Find this resource:

Bardou, I., Leprince, J., Chichery, R., Vaudry, H., & Agin, V. (2010). Vasopressin/oxytocin-related peptides influence long-term memory of a passive avoidance task in the cuttlefish, Sepia officinalis. Neurobiology, Learning, and Memory, 93, 240–247.Find this resource:

Bauknecht, P., & Jékely, G. (2015). Large-scale combinatorial deorphanization of Platynereis neuropeptide GPCRs. Cell Reproduction, 12, 684–693.Find this resource:

Bednářová, A., Kodrík, D., & Krishnan, N. (2013). Unique roles of glucagon and glucagon-like peptides: Parallels in understanding the functions of adipokinetic hormones in stress responses in insects. Comparative Biochemistry and Physiology A, 164, 91–100.Find this resource:

Beets, I., Janssen, T., Meelkop, E., Temmerman, L., Suetens, N., Rademakers, S., … Schoofs, L. (2012). Vasopressin/oxytocin-related signaling regulates gustatory associative learning in C. elegans. Science, 338, 543–545.Find this resource:

Blenau, W., & Baumann, A. (2001). Molecular and pharmacological properties of insect biogenic amine receptors: Lessons from Drosophila melanogaster and Apis mellifera. Archives of Insect Biochemistry and Physiology, 48, 13–38.Find this resource:

Blenau, W., Erber, J., & Baumann, A. (1998). Characterization of a dopamine D1 receptor from Apis mellifera: Cloning, functional expression, pharmacology, and mRNA localization in the brain. Journal of Neurochemistry, 70, 15–23.Find this resource:

Blenau, W., & Thamm, M. (2011). 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, 381–394.Find this resource:

Bravarenko, N. I., Korshunova, T. A., Malyshev, A. Y., & Balaban, P. M. (2003). Synaptic contact between mechanosensory neuron and withdrawal interneuron in terrestrial snail is mediated by L-glutamate-like transmitter. Neuroscience Letters, 341, 237–240.Find this resource:

Brezina, V., Bank, B., Cropper, E. C., Rosen S., Vilim F. S., Kupfermann, I., & Weiss, K. R. (1995). Nine members of the myomodulin family of peptide cotransmitters at the B16-ARC neuromuscular junction of Aplysia. Journal of Neurophysiology, 74, 54–72.Find this resource:

Brown, E. R., & Piscopo, S. (2013). Synaptic plasticity in cephalopods; more than just learning and memory? Invertebrate Neuroscience, 13, 35–44. doi:10.1007/s10158-013-0150-4.Find this resource:

Brunelli, G., Spano, P., Barlati, S., Guarneri, B., Barbon, A., Bresciani, R., & Pizzi, M. (2005). Glutamatergic reinnervation through peripheral nerve graft dictates assembly of glutamatergic synapses at rat skeletal muscle. Proceedings of the National Academy of Sciences USA, 102, 8752.Find this resource:

Burnstock, G. (2014). The concept of cotransmission: focus on ATP as a cotransmitter and its significance in health and disease. European Review 22, 1–17.Find this resource:

Burnstock, G., & Verkhratsky, A. (2009). Evolutionary origins of the purinergic signalling system. Acta Physiologica, 195, 415–447.Find this resource:

Byrne, J. H., & Hawkins, R. D. (2015). Nonassociative learning in invertebrates. Cold Spring Harbor Perspectives in Biology, 7, a021675.Find this resource:

Cardoso, J. C. R., & Larhammar, D. (2014). Comparative evolution of peptide hormone-binding GPCRs: A route to understanding functional complexity. General and Comparative Endocrinology, 209, 1–2.Find this resource:

Carlson, S. L., Kempsell, A. T., & Fieber, L. A. (2012). Pharmacological evidence that D-Aspartate activates a current distinct from ionotropic glutamate receptor currents in Aplysia californica. Brain Behavior, 2, 391–401. doi: 10.1002/brb3.60Find this resource:

Cary, G. A., Cuttler, A. S., Duda, K. A., Kusema, E. T., Myers, J. A., & Tilden, A. R. (2011). Melatonin: Neuritogenesis and neuroprotective effects in crustacean x-organ cells. Comparative Biochemistry and Physiology A, 161, 355–360. doi:10.1016/j.cbpa.2011.12.005.Find this resource:

Caulfield, M. P., & Birdsall, N. J. (1998). International Union of Pharmacology. XVII. Classification of muscarinic acetylcholine receptors. Pharmacology Review, 50, 279–290.Find this resource:

Chang, E. S., Prestwich, G. D., & Bruce, M. J. (1990). Amino acid sequence of a peptide with both molt-inhibiting and hyperglycemic activities in the lobster, Homarus americanus. Biochemical and Biophysical Research Communications, 171, 818–826.Find this resource:

Christie, A. E., Stemmler, E. A., & Dickinson, P. S. (2010). Crustacean neuropeptides. Cellular and Molecular Life Sciences, 67, 4135–4169. doi: 10.1007/s00018-010-0482-8.Find this resource:

Chung, J. S., Dircksen, H., & Webster, S. G. (1999). A remarkable, precisely timed release of hyperglycemic hormone from endocrine cells in the gut is associated with ecdysis in the crab Carcinus maenas. Proceedings of the National Academy Sciences USA, 96, 13103–13107.Find this resource:

Chung, J. S., & Webster, S. G. (2003). Moult cycle-related changes in biological activity of moult-inhibiting hormone (MIH) and crustacean hyperglycaemic hormone (CHH) in the crab, Carcinus maenas. From target to transcript. European Journal of Biochemistry, 270, 3280–3288.Find this resource:

Croll, N. (1977). Sensory mechanisms in nematodes. Annual Review of Phytopathology, 15, 75–89.Find this resource:

D’aniello, A., Spinelli, P., De Simone, A., D’aniello, S., Branno, M., Aniello, F., … Rastogi, R. K. (2003). Occurrence and neuroendocrine role of D-aspartic acid and N-methyl- D-aspartic acid in Ciona intestinalis. FEBS Letters, 552, 193–198.Find this resource:

Davies, J. D., & Watkins, J. C. (1982a). Selective excitatory amino acid antagonist action of γ-D-glutamylaminomethyl-sulphonate (GAMS) on cat spinal neurones. Journal of Physiology London, 332, 108P–109P.Find this resource:

Davies, J. D., & Watkins, J. C. (1982b). Actions of D and L forms of 2-aminophosphonovalerate and 2-amino-4-phosphonobutyrate in the cat spinal cord. Brain Research, 235, 378–386.Find this resource:

Dickinson, P. S., Fairfield, W. P., Hetling, J. R., & Hauptman, J. (1997). Neurotransmitter interactions in the stomatogastric system of the spiny lobster: One peptide alters the response of a central pattern generator to a second peptide. Journal of Neurophysiology, 77, 599–610.Find this resource:

Dickinson, P. S., Mecsas, C., Hetling, J., & Terio, K. (1993). The neuropeptide red pigment concentrating hormone meets rhythmic pattern generation at multiple sites. Journal of Neurophysiology, 69, 1475–1483.Find this resource:

Dircksen, H., Böcking, D., Heyn, U., Mandel, C., Chung, J. S., Baggerman, G., … Webster, S. G. (2001). Crustacean hyperglycaemic hormone (CHH)-like peptides and CHH precursor-related peptides from pericardial organ neurosecretory cells in the shore crab, Carcinus maenas, are putatively spliced and modified products of multiple genes. Biochemistry Journal, 356, 159–170.Find this resource:

Donaldson, Z., & Young, L. (2008). Oxytocin, vasopressin, and the neurogenetics of sociality. Science, 322, 900–904.Find this resource:

Eccles, J. C. (1957). The clinical significance of research work on the chemical transmitter substances of the nervous system. Medical Journal of Australia, 44, 745–753.Find this resource:

Eccles, J. C. (1964). The physiology of synapses. Berlin, Germany: Springer Verlag.Find this resource:

Elekes, K., & Florey, E. (1987). Immunocytochemical evidence for the GABAergic innervation of the stretch receptor neurons in crayfish. Neuroscience, 22, 1111–1122.Find this resource:

Fingerman, M., Nagabhushanam, R., Sarojini, R., & Reddy, P. S. (1994). Biogenic amines in crustaceans: Identification, localization, and roles. Journal of Crustacean Biology, 14, 413–437.Find this resource:

Fujino, Y., Nagahama, T., Oumi, T., Ukena, K., Morishita, F., Furukawa, Y., … Nomoto, K. (1999). Possible functions of oxytocin/vasopressin-superfamily peptides in annelids with special reference to reproduction and osmoregulation. Journal of Experimental Zoology, 284, 401–406.Find this resource:

Fuse, T., Ikeda, I., Kita, T., Furutani, S., Nakajima, H., Matsuda, K., … Ozoe, Y. (2015). Synthesis of photoreactive ivermectin B-1a derivatives and their actions on Haemonchus and Bombyx glutamate-gated chloride channels. Pesticide Biochemistry and Physiology, 120, 82–90.Find this resource:

Futamachi, K. J. (1972). Acetylcholine: Possible neuromuscular transmitter in Crustacea. Science, 175, 1373–1375.Find this resource:

Gäde, G. (2009). Peptides of the adipokinetic hormone/red pigment-concentrating hormone family. Annals of the New York Academy of Science, 11631, 125–136.Find this resource:

Gäde, G., Simek, P., Clark, K., & Auerswald, L. (2006). Unique translational modification of an invertebrate neuropeptide: A phosphorylated member of the adipokinetic hormone peptide family. Biochemistry Journal, 393, 705–713.Find this resource:

Gallus, L., Ferrando, S., Bottaro, M., Girosi, L., Ramoino, P., Diaspro, A., … Tagliafierro, G. (2006). Distribution of choline acetyltransferase immunoreactivity in the alimentary tract of the barnacle Balanus amphitrite (Cirripedia, Crustacea). Neuroscience Letters, 409, 230–233.Find this resource:

Garrison, J. L, Macosko, E. Z., Bernstein, S., Pokala, N., Albrecht, D. R., & Bargmann, C. I. (2012). Oxytocin/vasopressin-related peptides have an ancient role in reproductive behavior. Science, 338, 540–543.Find this resource:

Gingl, E., French, A. S., Panek, I. L., Meisner, S., & Tokkeli, P. H. (2004). Dendritic excitability and localization of GABA-mediated inhibition of spider mechanoreceptor neurons. European Journal of Neuroscience, 20, 59–65.Find this resource:

Glowik, R. M., Golowasch, J., Keller, R., & Marder, E. (1997). D-glucose-sensitive neurosecretory cells of the crab Cancer borealis and negative feedback regulation of blood glucose level. Journal of Experimental Biology, 200, 1421–1431.Find this resource:

Goodson, J. (2008). Nonapeptides and the evolutionary patterning of sociality. Progress in Brain Research, 170, 3–15.Find this resource:

Grimmelikhuijzen, C. J. P., Williamson, M., & Hansen, G. N. (2002). Neuropeptides in cnidarians. Canadian Journal of Zoology, 80, 1690–1702. doi:10.1139/z02-137.Find this resource:

Gruber, C. (2014). Physiology of invertebrate oxytocin and vasopressin neuropeptides. Experimental Physiology, 99, 55–61.Find this resource:

Guan, Z-B., Shui, Y., Liao, X-R., Xu, Z-H., & Zhou, X. (2014). Primary structure of a novel gonadotropin-releasing hormone (GnRH) in the ovary of red swamp crayfish Procambarus clarkii. Aquaculture, 418, 67–71.Find this resource:

Hooper, S. L., O’Neil, M. B., Wagner, R., Ewer, J., Golowasch, J., & Marder, E. (1986). The innervation of the pyloric region of the crab, Cancer borealis: Homologous muscles in decapod species are differently innervated. Journal of Comparative Physiology A, 159, 227–240.Find this resource:

Inoue, I., Nagahama, T., & Takata, M. (1994). Cl- channel as a cholinergic ACh receptor responsible for generation of inhibitory junction potential in Aplysia buccal muscle cells. Japanese Journal of Physiology, 44, S149–S155.Find this resource:

Iwakoshi-Ukena, E., Ukena, K., Takuwa‐Kuroda, K., Kanda, A., Tsutsui, K., & Minakata, H. (2004). Expression and distribution of octopus gonadotropin-releasing hormone in the central nervous system and peripheral organs of the octopus (Octopus vulgaris) by in situ hybridization and immunohistochemistry. Journal of Comparative Neurology, 477, 310–323.Find this resource:

Jékely, G. (2013). Global view of the evolution and diversity of metazoan neuropeptide signaling. Proceedings of the National Academy of Sciences USA, 110, 8702.Find this resource:

Jin, X., Pokala, N., & Bargmann, C. I. (2016). Distinct circuits for the formation and retrieval of an imprinted olfactory memory. Cell, 164, 632–643.Find this resource:

Jo, Q., Laufer, H., Biggers, W., & Kang, H. (1999). Methyl farnesoate induced ovarian maturation in the spider crab, Libinia emarginata. Invertebrate Reproduction and Development, 36, 79–85.Find this resource:

Johnson, J., Kavanaugh, S., Nguyen, C., & Tsai, P. (2014). Localization and functional characterization of a novel adipokinetic hormone in the mollusk, Aplysia californica. PLoS ONE, 9.Find this resource:

Kabotyanski, E., Baxter, D., Cushman, S., & Byrne, J. (2000). Modulation of fictive feeding by dopamine and serotonin in Aplysia. Journal of Neurophysiology, 83, 374–392.Find this resource:

Kah, O., Lethimonier, C., Somoza, G., Guilgur, L. G., Vaillant, C., & Lareyre, J. J. (2007). GnRH and GnRH receptors in metazoa: A historical, comparative, and evolutive perspective. General and Comparative Endocrinology, 153, 346–364.Find this resource:

Kanda, A., Takahashi, T., Satake, H., & Minakata, H. (2006). Molecular and functional characterization of a novel gonadotropin- releasing-hormone receptor isolated from the common octopus (Octopus vulgaris). Biochemistry Journal, 395, 125–135.Find this resource:

Katusić, Z. S. (1992). Endothelial L-arginine pathway and regional cerebral arterial reactivity to vasopressin. American Journal of Physiology, 262, H1557–562.Find this resource:

Katz, P. S, & Frost, W. N. (1996). Intrinsic neuromodulation: altering neuronal circuits from within. Trends in Neuroscience, 19, 54–61.Find this resource:

Kauffman, A. S., & Rissman, E. F. (2004). The evolutionarily conserved gonadotropin releasing hormone II modifies food intake. Endocrinology, 145, 686–691.Find this resource:

Kawada, T., Kanda, A., Minakata, H., Matsushima, O., & Satake, H. (2004). Identification of a novel receptor for an invertebrate oxytocin/vasopressin superfamily peptide: Molecular and functional evolution of the oxytocin/vasopressin superfamily. Biochemistry Journal, 382, 231–237. 10.1042/BJ20040555.Find this resource:

Kehoe, J. (1972a). Ionic mechanisms of a two-component cholinergic inhibition of Aplysia neurons. Journal of Physiology, 225, 85–114.Find this resource:

Kehoe, J. (1972b). The physiological role of three acertylcholine receptors in synaptic transmission in Aplysia. Journal of Physiology, 225, 147–172.Find this resource:

Kehoe, J., & Vulfius, C. (2000). Independence of and interactions between GABA-, glutamate-, and acetylcholine-activated Cl conductances in Aplysia neurons. Journal of Neuroscience, 20, 8585–8596.Find this resource:

Kemenes, I., Kemenes, G., Andrew, R. J., Benjamin, P. R., & O’Shea, M. (2002). Critical time-window for NO-cGMP-dependent long-term memory formation after one-trial appetitive conditioning. Journal of Neuroscience, 22, 1414–1425.Find this resource:

Kempsell, A. T., & Fieber, L. A. (2015). Age-related deficits in synaptic plasticity rescued by activating PKA or PKC in sensory neurons of Aplysia californica. Frontiers in Aging and Neuroscience, 7, 173. doi: 10.3389/fnagi.2015.00173.Find this resource:

Korshunova, T. A., & Balaban, P. M. (2014). Nitric oxide is necessary for long-term facilitation of synaptic responses and for development of context memory in terrestrial snails. Neuroscience, 266, 127–135.Find this resource:

Kratsios, P., Stolfi, A., Levine, M., & Hobert, O. (2012). Coordinated regulation of cholinergic motor neuron traits through a conserved terminal selector gene. Nature Neuroscience, 15, 205.Find this resource:

Kravitz, E. (2000). Serotonin and aggression: Insights gained from a lobster model system and speculations on the role of amine neurons in a complex behavior. Journal of Comparative Physiology A, 186, 221–238.Find this resource:

Kristan, W. B., Calabrese, R. L., & Friesen, W. O. (2005). Neuronal control of leech behavior. Progress in Neurobiology, 76, 279–327.Find this resource:

Kulkarni, G. K., & Fingerman, M. (1986). Effects of two tranquilizers, reserpine and chlorpromazine, on neurosecretory cells and the ovary of the fiddler crab, Uca pugilator. General Pharmacology, 17, 671–683.Find this resource:

Lee, Y-S., Bailey, C. H., Kandel, E. R., & Kaang, B-K. (2008). Transcriptional regulation of long-term memory in the marine snail Aplysia. (Review). Molecular Brain, 1, 3.Find this resource:

Lee Johnson, J. (1978). The excitant amino acids glutamic and aspartic acid as transmitter candidates in the vertebrate central nervous system. Progress in Neurobiology, 10, 155–202.Find this resource:

Lindemans, M., Janssen, T., Beets, I. Temmerman, L., Meelkop, E., & Schoofs, L. (2011). Gonadotropin-releasing hormine and adipokinetic hormone signaling systems share a common evolutionary origin. Frontiers in Endocrinology, 2, 16.Find this resource:

Liu, Z., Zhou, Z., Wang, L., Dong, W., Qiu, L., & Song, L. (2016). The cholinergic immune regulation mediated by a novel muscarinic acetylcholine receptor through TNF pathway in oyster Crassostrea gigas. Developmental and Comparative Immunology, 65, 139–148.Find this resource:

Lloyd, P. E., & Willows, A. O. (1988). Multiple transmitter neurons in Tritonia. II. Control of gut motility. Journal of Neurobiology, 19, 55–67.Find this resource:

Marder, E. (1974). Acetylcholine as an excitatory neuromuscular transmitter in the stomatogastric system of the lobster. Nature, 251, 730–731.Find this resource:

Marder, E. (1976). Cholinergic motor neurones in the stomatogastric system of the lobster. Journal of Physiology, 257, 63–86.Find this resource:

MacDermott, A., Role, L., and Siegelbaum, S. (1999). Presynaptic ionotropic receptors and the control of transmitter release. Annual Review of Neuroscience, 22, 443–485.Find this resource:

Macginitie, G. (1934). The egg-laying activities of the sea hare, Tethys californicus (Cooper). Biological Bulletin, 67, 300–303.Find this resource:

Matsuda, K., Nakamura, K., Shimakura, S-I., Miura, T., Kageyama, H., Uchiyama, M., …Ando, H. (2008). Inhibitory effect of chicken gonadotropin-releasing hormone II on food intake in the goldfish, Carassius auratus. Hormones and Behavior, 54, 83–89.Find this resource:

Mayer, M., & Westbrook, G. (1987). Permeation and block of N-methyl-D-aspartic acid receptor channels by divalent cations in mouse cultured central neurones. Journal of Physiology, 394, 501–527.Find this resource:

Martínez-Padrón, M., Gray, W., & Lukowiak, K. (1992). Conopressin G, a molluscan vasopressin-like peptide, alters gill behaviors in Aplysia. Canadian Journal of Physiology and Pharmacology, 70, 259–267.Find this resource:

McEwen, B. B. (2004). General introduction to vasopressin and oxytocin: Structure/metabolism, evolutionary aspects, neural pathway/receptor distribution, and functional aspects relevant to memory processing. Advances in Pharmacology, 50, 1–50.Find this resource:

Meelkop, E., Temmerman, L., Schoofs, L., & Janssen, T. (2011). Signalling through pigment dispersing hormone-like peptides in invertebrates. Progress in Neurobiology, 93, 125–147.Find this resource:

Meyer-Rochow, V. B. (2001). The crustacean eye: Dark/light adaptation, polarization sensitivity, flicker fusion frequency, and photoreceptor damage. Zoological Sciences, 18, 1175–1197.Find this resource:

Milakovic, M., Ormerod, K. G., Klose, M. K., & Mercier, A. J. (2014). Mode of action of a Drosophila FMRFamide in inducing muscle contraction. Journal of Experimental Biology, 217, 1725–1736.Find this resource:

Minakata, H. (2010). Oxytocin/vasopressin and gonadotropin-releasing hormone from cephalopods to vertebrates. Annals of the New York Academy of Sciences, 1200, 33–42.Find this resource:

Moroz, L. L., Kocot, K. M., Citarella, M. R., Dosung, S., Norekian, T. P., Povolotskaya, I. S., … Kohn, A. B. (2014). The ctenophore genome and the evolutionary origins of neural systems. Nature, 510, 109–114. doi: 10.1038/nature13400.Find this resource:

Moroz, L. L., & Kohn, A. (2011). Parallel evolution of nitric oxide signaling: Diversity of synthesis and memory pathways. Frontiers in Bioscience, 16, 2008–2051.Find this resource:

Moroz, L., & Kohn, A. (2015). Unbiased view of synaptic and neuronal gene complement in Ctenophores: Are there pan-neuronal and pan-synaptic genes across Metazoa? Integrative and Comparative Biology, 55, 1028–1049.Find this resource:

Moroz, L., & Kohn, A. (2016). Independent origins of neurons and synapses: Insights from ctenophores. Philosophical Transactions of the Royal Society London B, 371, 1685.Find this resource:

Nagaraju, G. P. C. (2007). Is methyl farnesoate a crustacean hormone. Aquaculture, 26, 39–54.Find this resource:

Nakatsuji, T., Han, D. W., Jablonsky, M. J., Harville, S. R., Muccio, D. D., & Watson, R. D. (2006). Expression of crustacean (Callinectes sapidus) molt-inhibiting hormone in Escherichia coli: Characterization of the recombinant peptide and assessment of its effects on cellular signaling pathways in Y-organs. Molecular and Cellular Endocrinology, 253, 96–104.Find this resource:

Nakatsuji, T., & Sonobe, H. (2004). Regulation of ecdysteroid secretion from the Y-organ by molt-inhibiting hormone in the American crayfish, Procambarus clarkii. General and Comparative Endocrinology, 135, 358–364.Find this resource:

Nässel, D. (1996). Peptidergic neurohormonal control systems in invertebrates. Current Opinions in Neurobiology, 6, 842–850.Find this resource:

Nishino, A. R., Okamura, Y., Piscopo, S., & Brown, E. (2010). A glycine receptor is involved in the organization of swimming movements in an invertebrate chordate. BMC Neuroscience, 11, 6.Find this resource:

Nusbaum, M. P., & Marder, E. (1988). A neuronal role for a crustacean red pigment concentrating hormone-like peptide: Neuromodulation of the pyloric rhythm in the crab, Cancer borealis. Journal of Experimental Biology, 135, 165–181.Find this resource:

Oumi, T., Ukena, K., Matsushima, O., Ikeda, T., Fujita, T., Minakata, H., & Nomoto, K. (1996). Annetocin, an annelid oxytocin-related peptide, induces egg-laying behavior in the earthworm, Eisenia foetida. Journal of Experimental Zoology, 276, 151–156.Find this resource:

Passano, L. M. (1953). Neurosecretory control of molting in crabs by the X-organ sinus gland complex. Physiology and Comparative Oecologia, 3, 155–189.Find this resource:

Pennings, S. C. (1991). Reproductive behavior of Aplysia californica Cooper: Diel patterns, sexual roles and mating aggregations. Journal of Experimental Marine Biology and Ecology, 149, 249–266.Find this resource:

Pizzi, M., Brunelli, G., Barlati, S., & Spano, P. (2006). Glutamatergic innervation of rat skeletal muscle by supraspinal neurons: A new paradigm in spinal cord injury repair. Current Opinions in Neurobiology, 16, 323–328.Find this resource:

Purves, D., Augustine, G. J., Fitzpatrick, D., Katz, L. C., LaMantia, A-S., & McNamara, J. O. (Eds). (2001). Neuroscience (2nd ed.). Sunderland, MA: Sinauer Associates.Find this resource:

Ranga Rao, K., & Riehm, J. (1988). Pigment-dispersing hormones: A novel family of neuropeptides from arthropods. Peptides, 9, 153–159.Find this resource:

Rinholm, J. E., Slettaløkken, G., Marcaggi, P., Skare, Ø., Storm-Mathisen, J., & Bergersen, L. H. (2007). Subcellular localization of the glutamate transporters GLAST and GLT at the neuromuscular junction in rodents. Neuroscience, 145, 579.Find this resource:

Roch, G. J., Busby, E. R., & Sherwood, N. M. (2011). Evolution of GnRH: Diving deeper. General and Comparative Endocrinology, 171, 1–16.Find this resource:

Ryan, J. F., Pang, K., Schnitzler, C. E., Nguyen, A-D., Moreland, R. T., Simmons, D. K., … Baxevanis, A. D. (2013). The genome of the ctenophore Mnemiopsis leidyi and its implications for cell type evolution. Science, 342, 1336–1345.Find this resource:

Sakamoto, T., Ogawa, S., Nishiyama, Y., Akada, C., Takahashi, H., Watanabe, T., Sakamoto, H. (2015). Osmotic/ionic status of body fluids in the euryhaline cephalopod suggests possible parallel evolution of osmoregulation. Science Reports, 5, 14469.Find this resource:

Santama, N., Benjamin, P. R., & Burke, J. F. (1995). Alternative RNA splicing generates diversity of neuropeptide expression in the brain of the snail Lymnaea: In situ analysis of mutually exclusive transcripts of the FMRFamide gene. European Journal of Neuroscience, 7, 65–76.Find this resource:

Santos, E. A., & Keller, R. (1993). Regulation of circulating levels of the crustacean hyperglycemic hormone: Evidence for a dual feedback control system. Journal of Comparative Physiology A, 163, 374–379.Find this resource:

Santos, E. A., Nery, L. E. M., Keller, R., & Goncalves, A. A. (1997). Evidence for the involvement of the crustacean hyperglycemic hormone in the regulation of lipid metabolism. Physiology and Zoology, 70, 415–420.Find this resource:

Satake, H., Takuwa, K., Minakata, H., & Matsushima, O. (1999). Evidence for conservation of the vasopressin/oxytocin superfamily in Annelida. Journal of Biological Chemistry, 274, 5605–5611.Find this resource:

Sattelle, D. B., Jones, A. K., Sattelle, B. M., Matsuda, K., Reenan, R., & Biggin, P. C. (2005). Edit, cut and paste in the nicotinic acetylcholine receptor gene family of Drosophila melanogaster. BioEssays, 27, 366–376.Find this resource:

Sedlmeier, D. (1988). The crustacean hyperglycemic hormone (CHH) releases amylase from the crayfish midgut gland. Regulatory Peptides, 20, 91–98.Find this resource:

Shomrat, T., Turchetti-Maia, A., Stern-Mentch, L., Basil, N., & Hochner, J. (2015). The vertical lobe of cephalopods: An attractive brain structure for understanding the evolution of advanced learning and memory systems. Comparative Physiology A, 201, 947–956.Find this resource:

Siegelbaum, S. A., Camardo, J. S., & Kandel, E. R. (1982). Serotonin and cyclic AMP close single K+ channels in Aplysia sensory neurones. Nature, 299, 413–417.Find this resource:

Sossin, W., & Abrams, T. (2009). Evolutionary conservation of the signaling proteins upstream of cyclic AMP-dependent kinase and protein kinase C in gastropod mollusks. Brain Behavior and Evolution, 74, 191–205.Find this resource:

Stein, W. R., Smarandache, C. B., Nickmann, M., & Hedrich, U. (2006). Functional consequences of activity-dependent synaptic enhancement at a crustacean neuromuscular junction. Journal of Experimental Biology, 209, 1285–1300.Find this resource:

Strumwasser, F., Jacklet, J. W., & Alvarez, R. B. (1969). A seasonal rhythm in the neural extract induction of behavioral egg-laying in Aplysia. Comparative Biochemistry and Physiology, 29, 197–206.Find this resource:

Stuart, A. E., Borycz, J., & Meinertzhagen, I. A. (2007). The dynamics of signaling at the histaminergic photoreceptor synapse of arthropods. Progress in Neurobiology, 82, 202–227.Find this resource:

Sugahara, R., Tanaka, S., Jouraku, A., & Shiotsuki, T. (2016). Functional characterization of the corazonin-encoding gene in phase polyphenism of the migratory locust, Locusta migratoria (Orthoptera: Acrididae). Applied Entomology and Zoology, 51, 225–232.Find this resource:

Sullivan, J., Sandeman, D., Benton, J., & Beltz, B. (2007). Adult neurogenesis and cell cycle regulation in the crustacean olfactory pathway: From glial precursors to differentiated neurons. Journal of Molecular Histology, 38, 527–542.Find this resource:

Susswein, A. J., & Chiel, H. J. (2012). Nitric oxide as a regulator of behavior: new ideas from Aplysia feeding. Progress in Neurobiology, 97, 304–317.Find this resource:

Tanaka, S., Harano, K-I., Nishide, Y., & Sugahara, R. (2016). The mechanism controlling phenotypic plasticity of body color in the desert locust: Some recent progress. Current Opinions in Insect Science, 17, 10–15.Find this resource:

Temple, J. L., Millar, R. P., & Rissman, E. F. (2003). An evolutionarily conserved form of gonadotropin-releasing hormone coordinates energy and reproductive behavior. Endocrinology, 144, 13–19.Find this resource:

Tierney, A. (2001). Structure and function of invertebrate 5-HT receptors: A review. Comparative Biochemistry and Physiology A, 128, 791–804.Find this resource:

Trimmer, B. A. (1995). Current excitement from insect muscarinic receptors. Trends in Neuroscience, 18, 104–111.Find this resource:

Tsai, P. (2006). Gonadotropin-releasing hormone in invertebrates: Structure, function, and evolution. General and Comparative Endocrinology, 148, 48–53.Find this resource:

Tsai, P-S., Sun, B., Rochester, J. R., & Wayne, N. L. (2010). Gonadotropin-releasing hormone-like molecule is not an acute reproductive activator in the gastropod, Aplysia californica. General and Comparative Endocrinology, 166, 280–288.Find this resource:

Tsai, P., & Zhang, L. (2008). The emergence and loss of gonadotropin-releasing hormone in protostomes: Orthology, phylogeny, structure, and function. Biology of Reproduction, 79, 798–805.Find this resource:

Ukena, K., Iwakoshi-Ukena, E., & Hikosaka, A. (2008). Unique form and osmoregulatory function of a neurohypophysial hormone in a urochordate. Endocrinology, 149, 5254–561.Find this resource:

Ukena, K., Oumi, T., Matsushima, O., Ikeda, T., Fujita, T., Minakata, H., & Nomoto, K. (1995). Effects of annetocin, an oxytocin-related peptide isolated from the earthworm Eisenia foetida, and some putative neurotransmitters on gut motility of the earthworm. Journal of Experimental Zoology, 272, 184–193.Find this resource:

Usherwood, P. N. R., Duce, I. R., & Boden, P. (1984). Slowly-reversible block of glutamate receptor channel by venoms of the spiders, Argiope trifasciata and Araneus gemma. Journal of Physiology Paris, 79, 241–245.Find this resource:

Van Golen, F. A., Li, K. W., De Lange, R. P. J., Jespersen, S., & Geraerts, W. P. M. (1995). Mutually exclusive neuronal expression of peptides encoded by the FMRFamide gene underlies a differential control of copulation in Lymnaea. Journal of Biochemistry, 270, 28487–28493.Find this resource:

van Kesteren, R. E., Smit, A. B., De Lange, R. P., Kits, K. S., Van Golen, F. A., Van Der Schors, … Geraerts, W. P. (1995). Structural and functional evolution of the vasopressin/oxytocin superfamily: Vasopressin-related conopressin is the only member present in Lymnaea, and is involved in the control of sexual behavior. Journal of Neuroscience, 15, 5989–5998.Find this resource:

Veenstra, J. A. (1989). Isolation and structure of corazonin, a cardioactive peptide from the American cockroach. FEBS Letters, 250, 231–234.Find this resource:

Veenstra, J. A. (2009). Does corazonin signal nutritional stress in insects? Insect Biochemistry and Molecular Biology, 39, 755–762.Find this resource:

Verlinden, H., Vleugels, R., Marchal, E., Badisco, L., Pflüger, H-J., Blenau, W., & Broeck, J. V. (2010). The role of octopamine in locusts and other arthropods. Journal of Insect Physiology, 56, 854–867.Find this resource:

Vyas, S., & Bradford, H. F. (1987). Co-release of acetylcholine, glutamate and taurine from synaptosomes of Torpedo electric organ. Neuroscience Letters, 82, 58–64.Find this resource:

Wainwright, G., Webster, S. G., Wilkinson, M. C., Chung, J. S., & H. H. Rees. (1996). Structure and significance of mandibular organ-inhibiting hormone in the crab, Cancer pagurus. Involvement in multihormonal regulation of growth and reproduction. Journal of Biological Chemistry, 271, 12749–12754.Find this resource:

Watkins, J. C. (1981a). Pharmacology of excitatory amino acid receptors. In P. J. Roberts, J. Storm-Mathisen, & G. A. R. Johnston, Glutamate: Transmitter in the central nervous system (pp. 1–24). New York, NY: Wiley.Find this resource:

Watkins, J. C. (1981b). Pharmacology of excitatory amino acid transmitters. In F. V. De Feudis & P. Mandel, Amino acid neurotransmitters (pp. 205–212). New York, NY: Raven Press.Find this resource:

Watkins, J. C., & Evans, R. H. (1981). Excitatory amino acid transmitters. Annual Review of Pharmacoligy and Toxicology, 21, 165–204.Find this resource:

Webster, S. G., Keller, R., & Dircksen, H. (2012). The CHH-superfamily of multifunctional peptide hormones controlling crustacean metabolism, osmoregulation, moulting, and reproduction. General and Comparative Endocrinology, 75, 217–233.Find this resource:

Weiss, K. R., Brezina, V., Cropper, E. C., Hooper, S. L., Miller, M. W., Probst, W. E., … Kupfermann, I. (1992). Peptidergic co-transmission in Aplysia: Functional implications for rhythmic behaviors. Experientia, 48, 456–463.Find this resource:

Wójciak, P., Remlinger-Molenda, A., & Rybakowski, J. (2012). The role of oxytocin and vasopressin in central nervous system activity and mental disorders. Psychiatrica Polska, 46, 1043–1052.Find this resource:

Zeleny, C. (1905). Compensatory regulation. Journal of Experimental Zoology, 2, 1–102.Find this resource: