Modulation of neuromuscular synapses and contraction in Drosophila 3rd instar larvae

Abstract

Over the past four decades, Drosophila melanogaster has become an increasingly important model system for studying the modulation of chemical synapses and muscle contraction by cotransmitters and neurohormones. This review describes how advantages provided by Drosophila have been utilized to investigate synaptic modulation, and it discusses key findings from investigations of cotransmitters and neurohormones that act on body wall muscles of 3rd instar Drosophila larvae. These studies have contributed much to our understanding of how neuromuscular systems are modulated by neuropeptides and biogenic amines, but there are still gaps in relating these peripheral modulatory effects to behavior.

KEYWORDS::

• Drosophila
• synaptic modulation
• neuropeptides
• octopamine
• tyramine
• FMRFamide
• proctolin
• muscle contraction
• third instar larvae

Introduction

The activation or inhibition of any neural pathway, ultimately depends on communication at chemical synapses, where neurotransmitters are released from presynaptic terminals, bind to receptors in the postsynaptic membrane (Figure 1) and alter electrical activity of the postsynaptic cell. ‘Classical’ neurotransmitters, such as acetylcholine, GABA and glutamate, bind to ionotropic receptors, chemically gated ion channels that directly alter the cell’s transmembrane potential. Other chemical signals, notably biogenic amines and neuropeptides, bind to metabotropic receptors that alter activity indirectly, typically through second messenger systems. Peptides and biogenic amines can be released as co-transmitters from presynaptic terminals to alter the effectiveness of ionotropic receptor activation (e.g. Adams & O’Shea, 1983; Fung et al., 1994; Nusbaum, Blitz, Swensen, Wood, & Marder, 2001), or they can be secreted by neuroendocrine cells and carried through the circulation to synapses, where they can alter transmitter release, postsynaptic responsiveness or both (e.g. Christie, Stemmler, & Dickinson, 2010; Ewer, 2005; Kravitz et al., 1980). These effects, which alter the efficacy of synaptic transmission, are referred to as ‘modulation’, and the signaling molecules that elicit them are often referred to as ‘modulators’ or ‘neuromodulators’. Thus, the ‘classical’ view of chemical synaptic transmission, in which a neurotransmitter acts directly to either depolarize or hyperpolarize a postsynaptic cell, represents only one component of synaptic communication; intercellular signaling in vivo involves many more components.

Figure 1. Sites of modulation at arthropod neuromuscular synapses. Electrical impulses in the motor axons trigger release of the primary neurotransmitter, L-glutamate, from small synaptic vesicles. Effects of the primary transmitter are mediated by ionotropic receptors and can be modulated via GPCRs that are activated by co-transmitters and neurohormones.

Classical transmitters number less than 10, but the number of modulators is larger due to the vast number of neuropeptides. There are at least 100 neuropeptides in the human central nervous system (Kastin, 2013). Analysis of sequence data from 10 animal phyla revealed 6225 distinct neuropeptide precursor proteins, each of which could be cleaved into one or multiple bioactive peptides (Jekely, 2013). Transcriptome mining typically predicts 105–216 distinct peptides in any given arthropod species (e.g. Christie, 2014; Christie et al., 2015). A new database for insect neuropeptides includes approximately 4700 amino acid sequences, representing 50 neuropeptide families from over 400 insect species (Yeoh et al., 2017). In Drosophila melanogaster at least 42 genes encode neuropeptide precursor polypeptides that predict a total of 75 distinct neuropeptides (Nässel & Winther, 2010). Only a subset has been detected in larvae and adult flies by chromatography and/or mass spectrometry, and it is not clear how many neuropeptides are actually expressed in Drosophila. Many neuropeptides appear to have redundant physiological effects, so the number of functionally distinct neuropeptides may be lower than the number of predicted peptides.

This review focuses on modulation of neuromuscular synapses and, specifically, insights provided by work with Drosophila as a model system. More comprehensive reviews of neuropeptide research in Drosophila and other insects have been provided by others (e.g. Hewes & Taghert, 2001; Nässel & Winther, 2010; Taghert, 1999; Vanden, 2001; Wegener & Veenstra, 2015; Yeoh et al., 2017).

Drosophila as a model system

The use of molecular methods to map transmitter and hormonal systems throughout the Drosophila nervous system has been quite successful, and it is worth noting some ground-breaking contributions Drosophila research has provided to the study of neuromodulation using molecular techniques. The first sequence of an insect neuropeptide gene, the dFMRFa gene (Nambu, Murphy-Erdosh, Andrews, Feistner, & Scheller, 1988; Schneider & Taghert, 1988) and the first characterization of a neuropeptide receptor, the Drosophila tachykinin-like receptor (Li, Wolfgang, Wu, North, & Forte, 1991), were reported before the Drosophila genome was released in 2000. These findings led directly to the identification of other genes encoding neuropeptides and their receptors in Drosophila and other insects. Drosophila P[GAL4] enhancer trap lines and conditional reporter lines were used for staining neurons, particularly those expressing green fluorescent protein (GFP) or other markers in specific neuronal classes. When used in combination with immunochemical stains for amines and peptides, this technique has allowed researchers to map distributions of transmitters and modulators throughout the central and peripheral nervous systems of Drosophila larvae and adults (e.g. Nässel & Winther, 2010; Python & Stocker, 2002; Santos et al., 2007). When combined with confocal microscopy, this approach provides very detailed staining of individual neurons and their fine processes. Following rapid success in identifying aminergic and peptidergic ligands for G protein-coupled receptors (GPCRs) in Drosophila (Brody & Cravchik, 2000; Cazzamali & Grimmelikhuijzen, 2002; Hewes & Taghert, 2001; Johnson, Bohn, et al., 2003; Johnson, Garczynski, et al., 2003; Meeusen et al., 2002), mapping the distributions of the various ‘de-orphaned’ GPCRs in neurons and other target cells progressed more slowly, due to the failure of some studies to examine the entire CNS or to low sensitivity of immunocytochemical staining methods for GPCRs that can be expressed in low abundance (Nässel & Winther, 2010). More thorough descriptions of the ‘connectomes’ for transmitters and hormones remain as goals to be achieved.

The sections below describe work on modulation of neuromuscular synapses in Drosophila by aminergic and peptidergic co-transmitters and hormones. These studies have focused on body wall muscles of 3rd instar larvae, which contain eight abdominal segments with 30 muscles in each hemi-segment. Each muscle is a readily identifiable single cell, and the innervation for each has been well characterized (Hoang & Chiba, 2001; Keshishian, Broadie, Chiba, & Bate, 1996; Keshishian et al., 1993). Most studies have involved ventral longitudinal muscles 6, 7, 12 and 13, which are amenable to electrophysiological recording with sharp electrodes (Jan & Jan, 1976), loose patch electrodes (Kurdyak, Atwood, Stewart, & Wu, 1994; Renger, Ueda, Atwood, Govind, & Wu, 2000; Xing, & Wu, 2018) and suction electrodes (Ganetzky & Wu, 1982). Muscle contractions can also be recorded from these larvae, and care has been taken in devising extracellular salines that permit these physiological studies (Stewart, Atwood, Renger, Wang, & Wu, 1994).

The larvae contain four distinguishable types of synaptic bouton, including Types Ib (‘big’) and Is (‘small’), both of which contain primarily small, clear synaptic vesicles, Type II boutons, containing dense core vesicles, and Type III boutons, containing dense core vesicles and Drosophila insulin-like peptide (Dilp) (Atwood, Govind, & Wu, 1993; Gorczyca, Augart, & Budnik, 1993; Jia, Gorczyca, & Budnik, 1993; Keshishian et al., 1993). The classical transmitter released by the motor neurons is glutamate and is localized in the small, clear synaptic vesicles (Jan & Jan, 1976; Johansen, Halpern, Johansen, & Keshishian, 1989). The dense core vesicles contain substances that are thought to be released as co-transmitters, including octopamine (Monastirioti et al., 1995), pituitary adenylate cyclase-activating peptide (PACAP) (Zhong & Peña, 1995), proctolin (Anderson, Halpern, & Keshishian, 1988; Taylor et al., 2004), leukokinin (Cantera & Nässel, 1992) and Dilp (in Type III boutons; Gorczyca et al., 1993). The neuromuscular junctions and muscle fibers are also modulated by substances thought to be released as neurohormones, such as peptides encoded by the dFMRFa gene (Schneider, Roberts, & Taghert, 1993; Taghert, 1999) and proctolin (Taylor et al., 2004).

Synaptic boutons are amenable to electrical stimulation and recording with loose patch electrodes and to assessing calcium levels with fluorescent dyes. The terminal types exhibit physiological differences, with Type Ib motor neurons firing at lower impulse frequencies than do Type Is (Chouhan, Zhang, Zinsmaier, & Macleod, 2010). Type Ib terminals are larger than Type Is terminals and generate smaller EJPs that facilitate more strongly (Kurdyak et al., 1994; Lnenicka & Keshishian, 2000). Type II terminals release OA and other substances that modulate synaptic transmission at Type I synapses (see below). The three axon terminal types exhibit differences in calcium signaling, as revealed using calcium-sensitive fluorescent dyes while stimulating the motor axons (Xing & Wu, 2018). Type II terminals respond to low impulse frequencies, Type Is respond to intermediate frequencies and Type Ib respond only to high frequencies. The differences in calcium signaling stem from differences in expression of potassium channels and to different sodium channel alleles that affect excitability (Xing & Wu, 2018). Terminal Types Ib and II also differ in sensitivity to Ziram, a fungicide that increases excitability and generates calcium influx (Martin et al., 2016).

Octopamine (OA)

Octopamine has long been established as a neurotransmitter, neurohormone and neuromodulator in invertebrates, particularly arthropods, and is thought to function in ‘fight-or-flight’ responses, analogous to epinephrine and norepinephrine in vertebrates (reviewed by Evans & Maqueira, 2005; Evans & Robb, 1993; Farooqui, 2012; Roeder, 1999; Verlinden et al., 2010). In insects, OA plays key roles in desensitizing sensory inputs, initiating and maintaining rhythmic behaviors such as flight, and regulating complex social behaviors such as fighting, courtship and reproduction (e.g. Brembs, Christiansen, Pflüger, & Duch, 2007; Certel, Savella, Schlegel, & Kravitz, 2007; Dierick, 2008; Giurfa, 2006; Hana & Lange, 2017a, 2017b; Hoyer et al., 2008; Rezaval, Nojima, Neville, Lin, & Goodwin, 2014; Suver, Mamiya, & Dickinson, 2012; Wasserman et al., 2015). Most or all of these functions involve effects of OA within the CNS. At the peripheral level, OA’s actions as a neurohormone include mobilization of carbohydrates and lipids to help meet increased energy demands during increased activity and modulation of synaptic and contractile properties of muscles (Farooqui, 2012; Roeder, 1999; Verlinden et al., 2010).

The distribution of octopaminergic neurons in the central and peripheral nervous systems has been very well characterized in many insects, including Drosophila (Busch, Selcho, Ito, & Tanimoto, 2009; Farooqui, 2012; Monastirioti et al., 1995; Monastirioti, Linn, & White, 1996). Immunohistochemical staining in 3rd instar larvae revealed very similar staining patterns for OA and tyramine-beta-hydroxylase (TBH), which converts tyramine (TA) to OA (Monastirioti et al., 1995, 1996). Monastirioti et al. (1995) reported that OA is rarely observed in muscles 3–7, 25 and 28 but is usually present in the other 23 muscles. They also concluded that all Type II terminals they examined contained OA. This latter observation would imply the presence of OA in axons innervating all muscles except 6, 7 and 25–29, based on the innervation established by Hoang and Chiba (2001), who identified three axons that supply these 23 muscle fibers with Type II terminals. Closer inspection of the data from Monastirioti et al. (1995) indicates that OA is present less than 10% of the time in muscles 5, 6, 7, 25 and 28. In any case, most of the body wall muscles appear to receive octopaminergic innervation. Although only two octopaminergic axons appeared to project to the body wall muscles (Monastirioti et al., 1995), the appearance of OA on so many fibers would require its presence in at least three axons, probably the three giving rise to Type II terminals. OA is actually present in both dense core vesicles and small synaptic vesicles, due to the presence of different transport carriers on them (Grygoruk et al., 2014).

TA is present in all octopaminergic neurons as a precursor to OA, but it is not clear whether larval motor neurons contain TA but not OA. This issue could be addressed by comparing OA-staining to TA-staining using an anti-serum against TA that detects TA-like immunoreactivity in Drosophila (Nagaya, Kutsukake, Chigusa, & Komatsu, 2002), but there seems to have been little attempt to do so. Release of OA from Type II axon terminals has been quantified amperometrically using microelectrodes (Majdi et al., 2015). In that study, Type II terminals innervating muscles 12 and 13 were visualized using a fluorescent marker and were activated optogenetically using the light-activated ion channel, channelrhodopsin-2. The time-course of the amperometric currents was consistent with the opening and closing of a nanopore, suggesting ‘kiss-and-run’ release, and the quantal size was estimated at approximately 23,000 OA molecules per vesicle (Majdi et al., 2015). Pyakurel, Champaloux, and Venton (2016) used fast scan cyclic voltammetry to demonstrate release of OA from neurons in the ventral nerve cord of 3rd instar larvae. They expressed a red light-sensitive channel rhodopsin in neurons expressing tyrosine decarboxylase (TDC), which converts tyrosine to TA, the first step in synthesizing OA. Disulfiram, which inhibits TBH (Musacchio, Goldstein, Anagnoste, Poch, & Kopin, 1966), reduced the voltammetric signal by 80%, suggesting that the majority of material released was OA. Methods should be developed to detect TA in order to determine whether it is released from octopaminergic neurons or only from cells that contain TDC but not TBH.

Although an early report (Nishikawa & Kidokoro, 1999) suggested that OA decreases synaptic currents in larval muscles, subsequent studies (Koon et al., 2011; Nagaya et al., 2002; Ormerod, Hadden, Deady, Mercier, & Krans, 2013) have shown consistently that OA increases the amplitude of excitatory junctional potentials (EJPs). The increase in EJP amplitude is not accompanied by a change in amplitude of spontaneous miniature excitatory potentials (Koon et al., 2011), indicating that OA increases the number of quanta of transmitter released per nerve impulse. OA also elicits neuropeptide release from Type 1b boutons on muscles 6 and 7, even in the absence of extracellular Ca²⁺, through a cAMP-dependent process that liberates Ca²⁺ from internal stores via ryanodine receptors and inositol triphosphate (IP₃) receptors (Shakiryanova, Zettel, Gu, Hewes, & Levitan, 2011). Unlike OA, TA reduces EJP amplitude, and this effect is blocked by yohimbine, a TA receptor antagonist (Donini & Lange, 2004; Nagaya et al., 2002; Ormerod et al., 2013).

Receptors for OA and TA are expressed in larval body wall muscles, based on imaging using promoter Gal4 lines (El-Kholy et al., 2015). The longitudinal, transverse, oblique & acute body wall muscles in both dorsal & ventral regions all express the octopamine receptor Oct[beta]2, and a smaller number of lateral longitudinal, ventral longitudinal and lateral oblique muscles express the tyramine receptor TyrR III. Although no images of motor neuron terminals were reported, promoter regions for some receptors are expressed in the larval CNS and in some cases in segmental nerves. Their results suggest at least Oct[beta]1R, Oct[beta]3R, TyrR, TyrRII and TyrRIII as candidates for presynaptic receptors. Other work (Koon et al., 2011) has demonstrated expression of Oct[beta]2 and its role as an autoreceptor regulating neuritic outgrowth and synaptogenesis.

In addition to enhancing synaptic transmission in 3rd instar larvae, OA increases the amplitude of nerve-evoked contractions and induces contractions that can be even larger than those evoked synaptically (Ormerod et al., 2013). These effects are not blocked by yohimbine or mimicked by TA. OA-induced contractions probably reflect direct actions on muscle fibers, and the enhancement of nerve-evoked contractions probably reflects both presynaptic and postsynaptic actions, as reported for other arthropods (e.g. Evans, 1984; Hoyle, 1978; Kravitz et al., 1980). Surgical ablation of individual muscle cells indicates stronger effects of OA on larval muscles 12 and 13 than 6 and 7, and the same cell-selectivity is apparent for OA’s enhancement of EJPs (Ormerod et al., 2013). This cell-selectivity ‘correlates’ with the presence of OA in Type II boutons on cells 12 and 13 and its rare occurrence in boutons innervating cells 6 and 7 (Monastirioti et al., 1995). On the other hand, OA’s ability to induce contractions of cells 6 and 7 suggests the presence of OA receptors in these fibers. It would be interesting to assess levels of OA receptor expression in the four muscle cells and the motor neurons innervating them to determine the extent to which receptor expression in muscle depends on the presence of OA presynaptically.

Modulatory effects of OA play important roles in crawling behavior in 3rd instar larvae. Mutant larvae (t[beta]h^nM18) with impaired TBH function contain elevated TA levels and reduced OA levels. These larvae exhibit reduced crawling speed and spend more time pausing than wild type larvae, and these effects are rescued to some extent by feeding larvae OA or the TA receptor antagonist, yohimbine (Saraswati, Fox, Soll, & Wu, 2004). The tbh^nM18 mutants also show dramatic reductions in the rate of rhythmic contractions of the body segments, in the frequency of impulse bursts in the nerves supplying the body wall muscles, and in the rate of bursts of synaptic potentials in the muscles (Fox, Soll, & Wu, 2006). Thus, OA and TA play roles in initiating and modulating crawling rhythm.

At least one context in which OA modulates crawling is an increase in foraging behavior during starvation (Koon et al., 2011). Starvation elicits increased crawling speed, a long-lasting increase in transmitter release accompanied by increased EJP amplitude, and increased outgrowth and synaptogenesis of Type I and Type II nerve terminals. These effects are absent in tbh^nM18 mutants but are rescued by inserting a TBH transgene in just the octopaminergic neurons. Expressing a cell death protein (Hid) specifically in Type II neurons produces a phenotype very similar to tbh^nM18 larvae. Thus, during starvation release of OA from Type II boutons enhances their own arborization and that of nearby Type I boutons, and increases transmitter output and muscle contractions to support more rapid foraging. All these effects are mediated Oct[beta]2 receptors but not OAMB receptors, and they involve activation of CREB via cAMP (Koon et al., 2011). Experiments with dunce (dnc) and rutabega (rut) mutants, which alter phosphodiesterase activity and calcium-dependent activation of adenylyl cyclase, respectively, have also implicated cAMP in altering arborization of motor axon terminals and synaptic bouton morphology (Ueda & Wu, 2012; Zhong, Budnik, & Wu, 1992) and in altering nerve terminal excitability, transmitter release, synaptic facilitation and post-tetanic potentiation (Ueda & Wu, 2009; Zhong & Wu, 1991). cAMP appears to play a role in synaptic homeostasis by enhancing transmitter output to compensate for a loss of muscle fiber input resistance when fibers enlarge (Ueda & Wu, 2015).

Proctolin

Proctolin was the first insect neuropeptide identified (Brown & Starratt, 1975) and is broadly conserved across the arthropods (Christie, 2014, 2015; Christie et al., 2010; Isaac, Taylor, Hamasaka, Nässel, & Shirras, 2004; Orchard, Belanger, & Lange, 1989) but appears to be absent from a few insect species (Caers et al., 2012). Proctolin acts as a co-transmitter and neurohormone and modulates neuromuscular synapses, muscle contraction, cardiac contraction, circulation, stomach and gut motility, reproductive organs and sensory coding (Adams & O’Shea, 1983; Erxleben, deSantis, & Rathmayer, 1995; Kravitz et al., 1980; Lange, 2002; Marder, Hooper, & Siwicki, 1986; Orchard et al., 1989; Pasztor & MacMillan, 1990). The first gene encoding a proctolin precursor protein was identified in Drosophila (Taylor et al., 2004). Evidence for the presence of the proctolin peptide (RYLPT) in Drosophila is based on: (a) chromatographic analysis of CNS and tissue extracts with subsequent testing using bioassay and immunoassay (Anderson et al., 1988), (b) immunohistochemical staining using antibodies against proctolin or the proctolin precursor (Anderson et al., 1988; Taylor et al., 2004), (c) in situ hybridization with a riboprobe to the Proct gene (Taylor et al., 2004) and (d) the presence of proctolin in clones derived from a cell line of Drosophila CNS (Ui-Tei, Sakuma, Watanabe, Miyake, & Miyata, 1995). Extraction and sequencing of the authentic peptide from Drosophila, however, have not been achieved. We recently subjected an extract from ten Drosophila 3rd instar larvae to matrix-assisted laser desorption/ionization time of flight (MALDI-TOF) mass spectrometry, and our results (Figure 2) show a very small peak corresponding to a molecular mass (649.6 Da) close to that predicted for proctolin (648 Da). We do not know why recovery of substantial amounts of proctolin from Drosophila extracts is so difficult.

Figure 2. Matrix-assisted laser desorption/ionization time of flight (MALDI-TOF) mass spectrometry of extracts from 10 Drosophila 3rd instar larvae reveals a small peak (arrow) at 649.6 Da, close to the predicted molecular mass of Proctolin (648 Da).

Proctolin-like immunoreactivity in 3rd instar Drosophila larvae occurs in cell bodies in the CNS, in nerve endings on the hindgut and in nerve endings on body wall muscles of the abdominal segments, but not in thoracic segments (Anderson et al., 1988). Proctolin-immunoreactivity was observed reliably on muscle cells 12, 13 and 4 (up to 80% of cells examined, depending on abdominal segment), less reliably on cells 2, 3, 10, 14, 15 and 19, and was not observed on cells 6 or 7. Proctolin expression in nerve endings on muscles 12 and 13 but not 6 and 7 suggests its localization in dense core vesicles (Atwood et al., 1993), but it is not clear whether the endings containing proctolin are Type II, Type III or both (Jia et al., 1993). Similar expression patterns for proctolin and OA (i.e. in axon terminals on muscles 12 and 13 but not 6 and 7) poses questions about whether these two substances are co-localized in dense core vesicles, whether they exert similar physiological and/or behavioral effects, and whether they act synergistically. Nerve terminals in the ring gland, a neurohaemal organ, express the transcript of the proctolin gene (Proct) and show positive staining with proctolin-antibody (Taylor et al., 2004). Thus, proctolin could modulate the body wall muscles and their neuromuscular junctions either as a cotransmitter or as a neurohormone.

The first gene encoding a proctolin receptor in any invertebrate was identified in Drosophila independently by two groups (Egerod et al., 2003b; Johnson, Garczynski, et al., 2003). The receptor, a GPCR, was expressed in human embryonic kidney (HEK) cells or Chinese hamster ovary (CHO) cells that showed increased bioluminescence in response to proctolin with an EC₅₀ of 0.3–0.6 nM. The receptor is selective for proctolin over other insect neuropeptides but responds weakly to sex peptide at concentrations exceeding 1 µM (Johnson, Garczynski, et al., 2003). Northern blots of mRNA immunohistochemistry with an antibody against the receptor protein confirmed expression in the CNS, Malpighian tubules, heart and hindgut, but no expression was reported for larval muscles (Egerod et al., 2003b; Isaac et al., 2004; Johnson, Garczynski, et al., 2003).

Physiological effects of proctolin on body wall muscles of 3rd instar larvae (Ormerod et al., 2016) suggest proctolin receptor expression. Proctolin increases the amplitude of nerve-evoked contractions at low concentrations (EC₅₀ ∼ 2 nM) and induces contractions at much higher concentrations (EC₅₀ ∼ 900 nM). The latter effect was reduced by conditional knock-down using the Gal4-UAS system to express RNA-interference (RNAi) against the proctolin receptor in muscle cells. RNAi expression in neurons did not alter proctolin’s effect, suggesting a postsynaptic action but no presynaptic action, and proctolin did not alter muscle EJPs. Proctolin-induced contractions involve extracellular Ca²⁺ and intracellular Ca²⁺ stores and are antagonized by the L-type channel blocker, nifedipine and an inhibitor of store-operated calcium channels (Ormerod et al., 2016). Muscle ablation indicated that proctolin induces contractions more strongly in muscles 4, 12 and 13 than in 6 and 7, and proctolin also enhances nerve-evoked contractions more strongly in 4, 12 and 13 than in 6 and 7 (Ormerod et al., 2016). These cell-selective effects of proctolin suggest that its receptor may be expressed more strongly in some muscle fibers than others, but this possibility remains to be examined. Proctolin’s cell-selectivity is also similar to that of OA (Ormerod et al., 2013). In each case the modulator acts more strongly on muscles receiving innervation containing that same modulator.

The substantial difference in dose-dependence between proctolin’s ability to induce contractions and its enhancement of nerve-evoked contractions (Ormerod et al., 2016) might suggest that the two effects involve receptors with differing affinities for proctolin. Knock-down experiments with RNAi were only used to examine proctolin-induced contractions and should be repeated to confirm that modulation of nerve-evoked contractions involves the same receptor. This outcome seems likely since the EC₅₀ for enhancement of evoked contractions is closer to the EC₅₀ values for proctolin to elicit effects on cultured cells transfected with the Drosophila proctolin receptor (Egerod et al., 2003b; Johnson, Garczynski, et al., 2003). Other arthropods, however, have more than one proctolin receptor (e.g. Christie et al., 2015), so this possibility should be ruled out.

Increasing the frequency and number of stimuli per burst lowers the EC₅₀ and threshold for proctolin to enhance nerve-evoked contractions, indicating activity-dependent modulation (Ormerod et al., 2016). Since proctolin does not appear to act presynaptically, the drop in threshold might involve a postsynaptic change in physiological state. Increasing the frequency and number of impulses would release more glutamate, elicit greater postsynaptic depolarization and higher levels of intracellular calcium that, in turn, might alter second messengers in the muscle cells. Alternatively, the higher stimulus levels might enhance the release of cotransmitters (Belanger & Orchard, 1993; Shakiryanova, Tully, Hewes, Deitcher, & Levitan, 2005) that might enhance the effectiveness of proctolin. Such second order modulation or ‘metamodulation’, has been reported in other systems (e.g. Marder, O'Leary, & Shruti, 2014; Mesce, 2002). The drop in threshold and EC₅₀ for proctolin to enhance contractions when motor neuron activity increases also suggests that neurohormones may be more effective in targeting active muscles, particularly when the same substance is released as a cotransmitter at neuromuscular junctions. High frequency stimulation of locust ovipositor motor neurons, however, appears to release enough proctolin to saturate postsynaptic receptors and decrease responsiveness to bath-applied proctolin (Belanger & Orchard, 1993). Release of proctolin and possible interactions with other cotransmitters should be investigated.

Knocking down proctolin receptor expression in muscle cells did not alter locomotion speed, but receptor knock down in neurons reduced the ability of larvae to crawl faster at higher ambient temperatures (Ormerod et al., 2016). This suggests that proctolin may play some role in temperature sensation or in mediating behavioral responses to temperature change. Such effects, however, do not appear to involve effects of proctolin on body wall muscles.

FMRFamides

FMRFamides and FMRFamide-like peptides are involved in many physiological functions, including regulating cardiac performance, circulation, feeding, digestion, neuromuscular transmission and muscle contraction (e.g. Greenberg & Price, 1992; Lange, 2001; Mercier, Friedrich, & Boldt, 2003). In Drosophila, they are implicated in cardiac function, locomotion, flight, ecdysis and stress-induced sleep (Agrawal, Sadaf, & Hasan, 2013; Kim et al., 2006; Klose, Dason, Atwood, Boulianne, & Mercier, 2010; Lenz, Xiong, Nelson, Raizen, & Williams, 2015; Maynard et al., 2013; Taghert, 1999). The dFMRFa gene (Nambu et al., 1988; Schneider & Taghert, 1988) encodes eight distinct peptides, five of which contain the amidated C-terminal sequence FMRFamide and three with sequences similar to FMRFamide (SAPQDFVRSamide, MDSNFIRFamide and SVQDNFMHFamide). The gene contains five copies of DPKQDFMRFamide, two copies of TPAEDFMRFamide and one copy of each of the other six peptides. Since dFMRFa is not expressed in motor neurons that project to larval body wall muscles but is expressed in Tv neuroendocrine neurons that project to neurohemal sites (Schneider et al., 1993; Taghert, 1999), the peptides are thought to modulate neuromuscular synapses and muscles as hormones, not cotransmitters.

Hewes, Snowdeal, Saitoe, and Taghert (1998) performed a detailed analysis of effects of the peptides encoded in dFMRFa on nerve-evoked contractions of larval body wall muscles. SAPQDFVRSamide had no effect, but the other seven peptides increased contraction amplitude, with thresholds of ∼10 nM for DPKQDFMRFamide, SPKQDFMRFamide, SDNFMRFamide, PDNFMRFamide and SVQDNFMHRamide, and 10-fold lower for TPAEDFMRFamide and MDSNFIRFamide. EC₅₀ values were ∼40 nM for most of the peptides, and their effects were equivalent whether applied separately or in combination, suggesting that the peptides are functionally redundant (Hewes et al., 1998). This view was supported by subsequent work identifying one GPCR that, when expressed in CHO cells and assayed for bioluminescence of co-transfected aequorin, fails to respond to SAPQDFVRSamide but responds to the other peptides encoded in dFMRFa with similar EC₅₀ values, ranging as low as 0.9–2 nM (Cazzamali & Grimmelikhuijzen, 2002; Meeusen et al., 2002). This GPCR also responded to Drosophila myosuppressin, sulfakinin-1 and short neuropeptide F, but at concentrations too high to be considered physiologically relevant (EC₅₀ values 38–110 nM), so it was designated the Drosophila FMRFamide receptor (FR). Genes for two receptors for Drosophila myosuppressin (DmsR-1 and DmsR-2) were subsequently identified, and when expressed in CHO cells and examined with a bioluminescence assay, they failed to respond to FMRFamide, Drosophila short neuropeptide F-1 and perisulfakinin (Egerod et al., 2003a). DmsR-2 was also expressed in HEK cells and assayed for translocation of [beta]-arrestin2-green fluorescent protein ([beta]ARR2-GFP), a protein involved in desensitization of nearly all GPCRs (Kohout & Lefkowitz, 2003). These cells responded to both Drosophila myosuppressin and DPKQDFMRFamide if a G-protein coupled receptor kinase was co-expressed to accelerate [beta]ARR2-GFP translocation (Johnson, Bohn, et al., 2003). HEK cells expressing either DmsR-1 or DmsR-2 also showed decreased cAMP levels in response to both myosuppressin and DPKQDFMRFamide but were ∼10-fold less sensitive to the latter peptide. Thus, many people have favoured the view that there is only one receptor for the peptides encoded in dFMRFa, at least at physiologically relevant peptide concentrations. Receptor and ligand modeling indicates that the five Drosophila peptides containing the sequence ‘FMRFamide’ exhibit subtle differences in docking and linking with the FMRFamide receptor, but they all elicit very similar responses in cardiac bioassays (Maynard et al., 2013).

Hewes et al. (1998) also found that DPKQDFMRFamide increases synaptic current in muscle fiber 6, indicating increased transmitter release from presynaptic axons. Dunn and Mercier (2005) showed that this peptide increases EJP amplitude in fibers 6 and 7 when the motor axon with Type 1b terminals is activated but not when Type Is terminals are activated, indicating a neuron-specific peptide effect. Using quantal current recordings, Klose et al. (2010) confirmed that DPKQDFMRFamide increases transmitter release from 1b terminals, and this effect was disrupted in larvae with mutations in FR and DmsR-2, implicating both receptors in mediating effects of DPKQDFMRFamide on transmitter output. Mutant FR and DmsR-2 lines also showed reduced escape responses to intense light, while mutant DmsR-1 larvae behaved normally. Escape responses were also reduced when RNAi was used in combination with the Gal4/UAS system to knock down FR and DmsR-2 expression in neurons, but responses were normal when knock down occurred in muscles (Klose et al., 2010). Finally, replacing larval hemolymph with physiological saline decreased fictive locomotor movement to near zero, suggesting a role for circulating hormones in maintaining these movements. Including DPKQDFMRFamide in the saline, however, maintained fictive locomotion, but the peptide’s effect was absent in larvae in which FR and DmsR-2 expression was knocked down in neurons with RNAi (Klose et al., 2010). Together, these results suggest that DPKQDFMRFamide acts as a hormone to help maintain larval locomotion, and that it acts specifically on neurons, probably by enhancing transmitter release from Type 1b terminals onto muscles. These effects depend on two GPCRs rather than just one. The involvement of more than one receptor offers potential for different peptides in the dFMRFa gene to elicit different physiological responses in different cells if expression levels of the receptors are not identical (Nusbaum & Blitz, 2012), but there is no direct evidence that the peptides act differently.

Molecular and pharmacological tools have been used to examine the intracellular signaling pathways underlying effects of DPKQDFMRFamide on larval motor neurons. The peptide’s ability to increase EJP amplitude is blocked by an inhibitor of calcium/calmodulin-dependent kinase II (CaMKII) and by conditional inhibition of CaMKII in transgenic larvae (ala1; Griffith et al., 1993) that express a CaMKII autoinhibitory peptide in response to heat shock (Dunn & Mercier, 2005). DPKQDFMRFamide increases the peak intracellular calcium concentration in synaptic boutons following single axonal stimuli, and this effect was abolished by depleting intracellular calcium stores using mutant larvae (CaP60A^Kum170) whose sarcoplasmic/endoplasmic reticulum calcium ATPase (SERCA) activity can be shut down by heat shock (Klose et al., 2010). The rise in intracellular calcium was also blocked by inhibitors of ryanodine receptors and IP₃ receptors, and by genetic disruption of these receptors, as indicated by a lack the peptide’s effect in mutants (Klose et al., 2010). As expected, inhibiting CaMKII did not block the peptide’s ability to increase intracellular calcium. Thus, in the presence of DPKQDFMRFamide, a nerve impulse generates calcium influx in the synaptic terminals which, in turn, appears to trigger release of calcium from intracellular stores via IP₃ receptors and ryanodine receptors. The elevated peak calcium concentration probably activates CaMKII, which phosphorylates some substrate that increases the number of quanta of transmitter released per nerve impulse (Klose et al., 2010).

DPKQDFMRFamide also induces small muscle contractions (Clark, Milakovic, Cull, Klose, & Mercier, 2008), and this effect is reduced by knock down of FR but not by knock down of DmsR-1 or DmsR-2 (Milakovic, Ormerod, Klose, & Mercier, 2014). When the Gal4/UAS system was used to achieve conditional expression of RNAi against FR, knocking down receptor expression in muscle reduced the effect of DPKQDFMRFamide, but knock down in nerve cells did not. This approach confirmed a postsynaptic site of action for the peptide, circumventing problems of trying to rule out the possibility that the peptide might also modulate transmitter release from extant nerve endings. The intracellular signaling systems underlying this postsynaptic effect of DPKQDFMRFamide are not known, but they do not appear to include cAMP, cGMP, IP₃, CaMKII or arachidonic acid, based on data obtained using physiological, pharmacological, biochemical and genetic approaches (Milakovic et al., 2014). The peptide-induced contractions require extracellular calcium and are inhibited by nifedipine (Clark et al., 2008), suggesting the involvement of L-type calcium channels in the sarcolemma (Gielow, Gu, & Singh, 1995). DPKQDFMRFamide also reduces input resistance in the muscle fibers, and this effect was also blocked by nifedipine (Ormerod, Krans, & Mercier, 2015). Thus, the peptide might activate L-type channels and generate calcium influx that, in turn, releases calcium from the sarcoplasmic reticulum to generate contractions, as occurs for other peptides acting on other insect muscles (Wegener & Nässel, 2000; Wilcox & Lange, 1995).

The ability of DPKQDFMRFamide to enhance nerve-evoked contractions is reduced by knocking down FR expression with RNAi (Ormerod et al., 2015). Reducing expression either in neurons or in muscle cells significantly decreased the peptide’s effect, and ubiquitous knock down decreased the effect even further. Thus, the peptide modulates muscle contraction through both presynaptic mechanisms that increase transmitter release and postsynaptic mechanisms that enhance the ability of the neurotransmitter (L-glutamate) to elicit contractions. Surgical ablation of muscle cells demonstrated that DPKQDFMRFamide enhances nerve-evoked contractions more strongly in fibers 6 and 7 than in 12 and 13, and the same cell-selectivity was observed for the peptide’s ability to induce contractions (Ormerod et al., 2015). In situ hybridization using fluorescently labeled probes for FR indicated higher expression of the receptor in fibers 6 and 7 than 12 and 13 (Ormerod et al., 2015). Thus, differences in the responsiveness of individual muscle cells to DPKQDFMRFamide appear to result from differences in expression levels of FR. Interestingly, the pattern of muscle cell-selectivity for modulation by DPKQDFMRFamide differs from and is complementary to the patterns reported for octopamine (Ormerod et al., 2013) and proctolin (Ormerod et al., 2016). DPKQDFMRFamide modulates contractions of fibers 6 and 7 more strongly than fibers 12 and 13, while proctolin and octopamine elicit stronger effects on 12 and 13 than on 6 and 7. The physiological and behavioral consequences of these differences in modulation are not known. We are still missing details that would provide a more complete picture of how larval body wall muscles are modulated, such as expression levels of the proctolin receptor and octopamine receptors in the muscle fibers and of FR and DmsR-2 in axon terminals.

Other cotransmitters

Less is known about neuromuscular modulation by other peptides in the motor axons supplying larval body wall muscles, although PACAP has received considerable attention. PACAP is a member of the secretin family of peptides and occurs in a 38 amino acid form (PACAP-38) and a 27 amino acid form (PACAP-27) (Arimura, 1992; Cardoso, Vieira, Gomes, & Power, 2010). A PACAP-like peptide is encoded in the Drosophila amnesiac (amn) gene (Feany & Quinn, 1995), but the sequence has not been identified. Immunoreactivity, detected with antibodies to vertebrate PACAP-38, is present in nerve terminals on muscle cells 4, 6, 7, 12 and 13 and is recoverable from fly extracts on Western blots (Zhong & Peña, 1995). Brief application of PACAP-38 depolarizes muscle fibers by up to 15 mV for tens of seconds, with a gradual repolarization phase. The depolarization is associated with inward Ca²⁺ current, and the repolarization is associated with outward K⁺ current. Stimulating the motor axons at 10 Hz mimics the effects of PACAP38, a slow depolarization with superimposed EJPs. The effects of stimulation are attenuated by desensitizing the receptors with prior exposure to the peptide (Zhong & Peña, 1995). Activation of the K⁺ current requires the Drosophila homolog of neurofibromatosis 1 (NF1) tumor suppressor protein, which activates the rut encoded adenylyl cyclase (Guo, The, Hannan, Bernards, & Zhong, 1997). Mutations in amn decrease L-type Ca²⁺-current in larval muscles, and the current is rescued by conditional expression of a wild type copy of amn or bath application of PACAP-38 (Bhattacharya, Lakhman, & Singh, 2004). Rescue by PACAP-38 is prevented by inhibiting adenylyl cyclase or by a Type 1 PACAP receptor antagonist (Bhattacharya et al., 2004). A Drosophila GPCR relatively selective for pigment-dispersing factor (PDF) also responds to calcitonin-like peptides and to PACAP (Mertens et al., 2005), but it is not clear whether this receptor mediates the effects of PACAP-38 on larval muscles. These observations suggest that a PACAP-like peptide acts as a cotransmitter to mediate slow depolarization and slow contractions in body wall muscles. PACAP-38 has been studied under conditions that avoid contractions, and its effects on contraction have not been recorded. In Drosophila amn plays important roles in memory formation and consolidation (DeZazzo, Xia, Christensen, Velinzon, & Tully, 1999; Keene et al., 2004), onset and maintenance of sleep (Liu, Guo, Lu, & Guo, 2008) and neural development (Hashimoto, Shintani, & Baba, 2002).

Drosophila genes encode eight Dilps, which play roles in life span, reproduction, development, organ growth, diapause and metabolism (Garelli et al., 2015; Nässel & Winther, 2010). Dilps 1–7 act through one receptor, DInr, and Dilp 8 acts through the neuronal relaxin receptor, Lgr3 (Brogiolo et al., 2001; Garelli et al., 2015). In larvae, Dilps 2, 3 and 5 are expressed in cells projecting to neurohemal areas, Dilp 7 is expressed in the abodominal nerve cord, and GFP-labeled Dilp 2 can be released from terminals of motor axons (Brogiolo et al., 2001; Wong et al., 2012). Dilps are present in axon terminals on muscle 12 and DInr is expressed in the muscle fibers (Gorczyca et al., 1993), but no modulatory effects of Dilps on larval synapses have been reported. In adults, diet-induced increases in insulin signaling increase expression of the synaptic protein, Complexin, which, in turn, reduces transmitter release from motor neurons onto CM9 muscles of the proboscis (Mahoney, Azpurua, & Eaton, 2016). Investigating similar effects in larvae might be fruitful.

Leucokinin (LK), a member of the kinin peptide family, is encoded in one Drosophila gene (Terhzaz et al., 1999) and is expressed in larvae in abdominal (ABLK) neurons that send axon terminals to muscle 8 (de Haro et al., 2010; Landgraf, Sánchez-Soriano, Technau, Urban, & Prokop, 2003). These terminals contain dense core vesicles, lie near spiracles and are not far from Malpighian tubules. This location might allow LK to act as a co-transmitter on muscle 8 or elicit paracrine effects on spiracles or Malpighian tubules. The ABLK cells also express serotonin receptors that mediate serotonin’s ability to suppress rearing and reduce turning during larval crawling (Okusawa, Kohsaka, & Nose, 2014). Possible effects of LK on muscle warrant further study.

Summary

Studies of the body wall muscles of Drosophila 3rd instar larvae have expanded our understanding of the roles of biogenic amines and peptides in synaptic modulation and behavior. This system allows researchers to combine genetic, pharmacological, biochemical, physiological and behavioral approaches to examine effects of cotransmitters and neurohormones on identified neurons and muscle cells. Conditional knock-down experiments are used effectively to distinguish presynaptic and postsynaptic effects. GPCRs for many modulators have been identified and are shown to work presynaptically and postsynaptically to alter transmitter release and responsiveness of postsynaptic cells to classical transmitters, and some intracellular mechanisms for these effects are identified. Modulators can act selectively on specific neurons and muscle cells, but the behavioral implications of these actions are not completely understood. Some modulators, notably PACAP, Dilps and LK, should be studied in greater detail.

Disclosure statement

No potential conflict of interest was reported by the authors.

Additional information

Funding

This study was supported by Natural Sciences and Engineering Research Council of Canada grant 46292.