Summary and Keywords
Nociception is a protective mechanism that mediates behavioral responses to a range of potentially damaging stimuli, including noxious temperature, chemicals, and mechanical stimulation. Nociceptive mechanisms are found throughout metazoans. Noxious stimuli are transduced by specialized, high-threshold peripheral nociceptors, which fire action potentials to elicit adaptive behavioral responses. Nociception is essential for survival and provides a mechanism for sensory perception of noxious stimuli, which alerts the organism to potential environmental dangers. When coupled with pain sensation and complex behavioral responses, this mechanism protects the organism from incipient damage. Moreover, acute and chronic pain may manifest as altered nociception in neuropathic pain states. Elucidating the neural bases of nociception is therefore important for identifying and implementing novel strategies for the treatment of neuropathic pain, as well as uncovering the mechanistic bases by which the nervous system integrates information to produce specific behaviors in response to a range of noxious stimuli. Invertebrate organisms, such as Drosophila melanogaster and Caenorhabditis elegans, have emerged as powerful, genetically tractable platforms for exploring these questions. Here, we concisely review the current state of knowledge regarding the cells, molecules, neural circuits, and behaviors associated with invertebrate nociception in the fruit fly and nematode worm.
Keywords: nociception, invertebrate, Drosophila, C. elegans, thermosensation, mechanosensation, chemosensation, sensitization, transient receptor potential (TRP) channels, Degenerin/epithelial sodium channel (DEG/ENaC)
Nociception is a process that is found throughout metazoans, which alerts organisms to noxious stimuli that are potentially tissue damaging or lethal under prolonged exposure. Transduction of noxious stimuli—such as mechanical, thermal, or chemical insults—occurs in primary sensory neurons known as nociceptors, which subsequently elicit distinct behavioral responses that trigger escape or other protective responses, allowing an animal to avoid incipient injury (Sherrington, 1906). Nociceptors are commonly high-threshold sensory neurons that exhibit free nerve endings (naked dendritic terminals) and may be myelinated (e.g., mammalian Αδ fibers) or unmyelinated (e.g., mammalian C fibers); these neurons function as cutaneous receptors for detecting noxious stimuli. Nociceptors are also distinguished from other sensory neurons with respect to sensitization following an intense stimulus or tissue damage. Nociceptive sensitization can manifest as hyperalgesia, involving an exaggerated response to a noxious stimulus, which may be due to nociceptor damage; or as allodynia, wherein animals exhibit nociceptive responses to a stimulus that is normally subthreshold, which may be caused by tissue damage leading to hypersensitization (Sandkühler, 2009). Studies across phyla including Chordata, Mollusca, Annelida, Nematoda, and Arthopoda have revealed conserved and distinct features of nociceptive biology, including molecular factors and signaling mechanisms, highlighting the utility of investigations in both vertebrates and invertebrates in uncovering potential mechanisms underlying human nociception and pain.
There is incredible diversity among invertebrates, and nociception has been studied in a variety of taxa, including Annelida, Arthropoda, Mollusca, and Nematoda. Herein, our discussion focuses on the nociceptive biology of the genetically tractable organisms Drosophila melanogaster and Caenorhabditis elegans—the most extensively studied of the invertebrates. Evidence of nociception in other invertebrates has been reviewed by Crook and Walters (2011) and Sneddon (2015).
Drosophila have a number of mechanosensory systems that allow them to sense auditory signals, monitor mechanical stress in the heart, and sense touches to the body (Ishikawa & Kamikouchi, 2016; Kernan, Cowan, & Zuker, 1994; Sénatore, Rami Reddy, Sémériva, Perrin, & Lalevée, 2010). Mechanosensory stimuli to the body can range from gentle and innocuous (albeit sometimes aversive) to noxious and potentially damaging; it is therefore important for animals to be able to distinguish between such events so that they can respond in a relevant fashion. Similarly, C. elegans has been useful in studying the neural basis of touch, as it was the first organism in which touch-insensitive animals were generated (Bounoutas & Chalfie, 2007).
Behaviorally, Drosophila third instar larvae challenged with noxious mechanical stimuli (≥45 mN Von Frey filament stimulation) perform a distinctive nocifensive escape behavior characterized by a 360˚ body-rolling response along the anterior-posterior body axis, wherein the animal rolls toward the direction of the stimulus (Tracey Jr., Wilson, Laurent, & Benzer, 2003; Hwang et al., 2007). This rolling behavior may have evolved as a means to defend against parasitoid wasps, which forcefully inject their ovipositor through the larval cuticle to lay their eggs (Carton & Frey, 1984). Rolling toward an attacking wasp serves to wrap the wasp ovipositor around the larva, disrupting the egg-laying process and allowing larval escape (Hwang et al., 2007).
In Drosophila larvae, studies have implicated peripheral nervous system Type II multidendritic (md) sensory neurons in mechanical and thermal nociception (Tracey Jr. et al., 2003; Hwang et al., 2007). Among the md neuron subtypes are the dendritic arborization (da) sensory neurons, which are subdivided into four morphologically distinct groups, referred to as classes I through IV (CI–CIV). CI neurons have the simplest dendritic architecture and CIV the most complex, the latter being characterized by space-filling arbors that exhibit dendritic tiling of the larval body wall (Grueber, Jan, & Jan, 2002; Grueber, Ye, Moore, Jan, & Jan, 2003). Here, da neurons have characteristic naked dendritic projections to the epidermis and are unmyelinated, similar to vertebrate C fiber nociceptors. Silencing synaptic transmission specifically in CIV neurons has been shown to strongly inhibit nocifensive rolling behavior to both noxious mechanical and thermal stimuli. Moreover, optogenetic activation of CIV neurons is sufficient to elicit this behavior, implicating CIV neurons as polymodal nociceptors (Hwang et al., 2007).
At the circuit level, larval nocifensive rolling is initiated in CIV nociceptive neurons, which synapse on multimodal Basin-2 and Basin-4 cells, two segmentally repeated projection neurons found in the sensory domain of the ventral nerve cord (VNC) (Ohyama et al., 2015; Vogelstein et al., 2014). The Basin neurons also receive synapses from vibration-sensitive chordotonal sensory neurons and are thus thought to be first-order interneurons for multimodal sensory integration (Ohyama et al., 2015). Network projections downstream from Basin interneurons follow a number of paths, some carrying Basin-subtype specific information and others carrying information integrated over many body segments. Second-order integration in this circuit occurs at commandlike Goro neurons, a pair of thoracic neurons projecting to the motor domain of the nerve cord that are also required for nocifensive rolling behavior (Ohyama et al., 2015). In addition to the Basin neurons, CIV neurons likely form synaptic contacts on A08n neurons, a class of ascending projection neurons that, when activated, promote nocifensive escape behavior (Vogelstein et al., 2014).
In C. elegans, brushing with an eyelash is the typical method of delivering a gentle touch stimulus to a worm—in order to deliver a harsh stimulus, platinum wire is pressed against the animal with a force of roughly 100–200 µN (Chatzigeorgiou et al., 2010; Li, Kang, Piggott, Feng, & Xu, 2011). Following a gentle touch, animals navigate away from the source of the stimulus, the distance moved generally being measured by the number of head swings (Goodman, 2006). A harsh touch elicits a similar response—anterior stimulation elicits backward locomotion, and posterior stimulation elicits forward locomotion—but over roughly three times the distance (Li et al., 2011). Touches to the anus of the animal elicit forward locomotion, while midbody touches initiate both backward and (more frequently) forward locomotion (Li et al., 2011).
At the cellular level, a variety of neuron subtypes have been implicated in C. elegans mechanosensation and nociception. PVD neurons, multidendritic sensory neurons covering the majority of the worm body wall, were first identified as gentle-touch sensitive (Way & Chalfie, 1989) and subsequently demonstrated to function as multimodal neurons with roles in sensing both noxious mechanical stimulation and acute cold shock (Chatzigeorgiou et al., 2010). PVD neurons were later found to be specifically involved in posterior harsh-touch sensation, along with PDE, a lateral, posterior sensory neuron with a sensory cilium (Li et al., 2011; White, Southgate, Thomson, & Brenner, 1986). ALM neurons are a set of nonciliated sensory neurons involved in both gentle and harsh posterior sensation via two molecular pathways (Chatzigeorgiou et al., 2010; Huang & Chalfie, 1994).
Via coablation experiments, several neurons have been implicated in anterior harsh-touch sensation: BDU, an interneuron that runs along the excretory canal; SDQR, an interneuron also responsible for high oxygen sensation; FLP neurons, multidendritic neurons covering the neck and head, also responsible for noxious heat sensation; ADE neurons, which are also associated with pheromone and gentle-touch sensation; and AQR, a neuron with immune and aerotaxic function, situated near the posterior pharynx and exposed to the pseudocoelomic fluid (Goodman, 2006; Li et al., 2011). PHA and PHB are two ciliated neurons, located in the phasmids (sensilla located just behind the rectum of the worm), which have been implicated in primary anal mechanical nociception. Ablation of PHA and PHB does not completely extinguish behavior, suggesting that other primary sensory neurons exist, with PVD and PDE potentially playing a minor role (Li et al., 2011). AVA and AVD are two of four pairs of interneurons that synapse on ventral nerve cord motor neurons and are necessary for backward locomotion commands following gentle touch (Chalfie et al., 1985). AVE neurons are additional backward command neurons that share synaptic targets with AVD neurons (Goodman, 2006). While AVA and AVD run the entire length of the ventral nerve cord, AVE runs only along the anterior portion of the cord (Goodman, 2006). When AVA, AVD, and AVE are ablated, animals fail to initiate backward locomotion following harsh touch, yet they retain forward locomotive responses (Li et al., 2011). PVC neurons, which are forward command neurons, have been found to be necessary for forward locomotion following posterior stimulation (Li et al., 2011). Forward locomotion facilitated by anal stimulation requires the PVC interneuron, as well as the DVA interneuron, which provides inputs to both anterior and posterior touch circuits (Goodman, 2006; Li et al., 2011; Wicks, Roehrig, & Rankin, 1996).
Molecular Regulation of Mechanical Nociception
As in vertebrates (Figure 1), several members of conserved ion channel families have been implicated in mechanical nociception in both Drosophila and C. elegans. Two transient receptor potential (TRP) channels, a group of nonselective cation channels, are known to be required for mechanical nociception in flies: Painless and Drosophila TRPA1 (dTRPA1) (Tracey Jr. et al., 2003; Zhong et al., 2012). Painless is expressed in larval md sensory neurons, including CIV, as well as chordotonal neurons, and mutations in painless increase the threshold of mechanical force required to initiate a nocifensive escape roll in larvae (Tracey Jr. et al., 2003). Painless shares protein domain similarity (Ankyrin repeats, or ARs) with the mechanically gated channel subunit NOMPC, another TRP channel that functions in gentle-touch mechanosensation (Árnadóttir & Chalfie, 2010; Bechstedt & Howard, 2008; Tracey Jr. et al., 2003; Yan et al., 2013). Mechanical gating, in response to gentle touch, is facilitated by ARs located at the N-terminus of NOMPC, which tether the channel to the microtubule cytoskeleton and allow touch-sensitive neuron activation (Zhang et al., 2015). The painless gene encodes multiple isoforms that vary in the number of N-terminal ARs. However, unlike NOMPC, these domains appear largely dispensable for Painless-mediated detection of noxious mechanical stimuli, as expression of a short isoform lacking most of the ARs can rescue painless mutant defects in mechanical, but not thermal, nociception (Hwang, Stearns, & Tracey, 2012). Likewise, dTRPA1 is required for mechanical nociception, where it is functionally required in CIV nociceptors; dTRPA1 encodes at least four distinct isoforms, including both thermosensory and nonthermosensory forms; and the dTRPA1-C/D isoforms exhibit specific expression in CIV nociceptors. While dTRPA1 genomic constructs can rescue mechanical nociception defects, expression of the nonthermosensory dTRPA1-C isoform in CIV neurons is insufficient to do so, indicating that an alternative isoform, perhaps dTRPA1-D, is required for mechanical nociception (Zhong et al., 2012).
At least two genes encoding subunits of the Degenerin/epithelial sodium channel (DEG/ENaC) protein family, a family of heterotrimeric channels permeable to sodium and calcium, are expressed and required in CIV nociceptors for mechanical nociception: pickpocket (ppk), and balboa/ppk-26 (Zhong, Hwang, & Tracey, 2010; Gorczyca et al., 2014; Guo, Wang, Wang, & Wang, 2014; Mauthner et al., 2014). Both genes encode very similar subunits that physically interact to form a functional heteromeric, voltage-insensitive ion channel required for mechanical, but not thermal, nociception (Gorczyca et al., 2014; Guo et al., 2014; Mauthner et al., 2014).
Piezo family proteins are evolutionarily conserved transmembrane proteins in Drosophila and mammals that are pore-forming subunits of mechanically gated channels (Coste et al., 2012). Drosophila Piezo (DmPiezo) exhibits expression in all types of sensory neurons, including CIV nociceptors, and is functionally required to permit mechanically activated inward currents in response to noxious stimuli (Kim, Coste, Chadha, Cook, & Patapoutian, 2012). It has been suggested that there are at least two distinct paths of noxious mechanical transduction in CIV neurons. Double mutant larvae for Dmpiezo and painless display a mechanical nociceptive defect similar to that seen when either gene is disrupted alone, suggesting that these molecules may function in a common pathway (Kim et al., 2012). In contrast, CIV-specific disruption of both Dmpiezo and ppk revealed a synergistic effect, resulting in the near-abolishment of mechanical nociception, suggesting that these molecules function in two parallel pathways within CIV neurons (Kim et al., 2012).
In C. elegans, harsh-touch mechanosensation in PVD and ALM neurons is regulated by expression of at least three genes—degt-1, mec-10, and mec-4—all which code for DEG/ENaC channel subunit proteins (Chatzigeorgiou et al., 2010; Huang & Chalfie, 1994; O’Hagan, Chalfie, & Goodman, 2005). Because expression of mec-10 alone does not lead to functional channel expression, mec-10 and mec-4 must be coexpressed (Bianchi et al., 2004; O’Hagan et al., 2005). The current hypothesis is that MEC-10 and DEGT-1 form a channel receptive to harsh touch, while MEC-10 and MEC-4 form a channel receptive to gentle touch (Chatzigeorgiou et al., 2010).
The nicotinic acetylcholine receptor (nAChR) subunits DEG-3, DES-2, and LGC-12 are expressed in nociceptive PVD neurons, and mutations in these subunits are linked to defects in PVD morphology, mechanosensory nociception, and cold detection. Furthermore, these nAChR subunits appear to enhance the response amplitude of PVD neurons to noxious stimuli, and mutants for these subunits exhibit reductions in cytosolic calcium levels that may underlie PVD morphological and behavioral defects (Cohen et al., 2014). The activity of PVD neurons can be further modulated by two proteins: the TRPM channel GTL-1 and the DEG/ENaC ASIC-1 (Husson et al., 2012). GTL-1 has been suggested to amplify the signal of PVDs, and ASIC-1 has been suggested to extend the dynamic range of PVDs—both appear to function downstream of primary mechanosensory channels (Husson et al., 2012).
Mechanosensation in PDE neurons, and thereby posterior harsh-touch sensation, are thought to be modulated via mechanical stimulation of PDE cilium, thereby activating the mechanically gated TRP-4 channel (Kang, Gao, Schafer, Xie, & Xu, 2010). TRP-4 is also expressed in DVA interneurons, which are involved in anal harsh-touch sensation; however, it does not appear to play the same role in these neurons (Li et al., 2011). The molecular mechanisms by which other mechanosensory neurons function in transducing harsh-touch stimuli in C. elegans are currently unknown.
Thermal Nociception and Sensitization
Larval and adult Drosophila both preferentially live at temperatures of approximately 24°C and are known to engage in a number of thermotactic behaviors that serve to maintain an optimum body temperature and allow the animals to avoid potentially dangerous thermal environments (Liu, Yermolaieva, Johnson, Abboud, & Welsh, 2003; Rosenzweig et al., 2005; Rosenzweig, Kang, & Garrity, 2008; Sayeed & Benzer, 1996). As with larval mechanical nociception, CIV md neurons function as high-temperature nociceptors, and third instar larvae respond to noxious heat (>42°C) in the same way that they do to harsh touch—with body rolling along the anterior-posterior axis (Tracey Jr. et al., 2003). The molecular underpinnings of heat- versus mechanical-evoked nocifensive responses are both conserved and distinct between nociceptive modalities. While the acquisition of rolling behavior in response to noxious heat does not occur before the third instar stage, optogenetic activation of CIV neurons elicits an aversive behavioral response, suggesting that CIV nociceptive function is present from the onset of larval development (Sulkowski, Kurosawa, & Cox, 2011).
Noxious cold stimuli elicit a number of behaviors in the Drosophila larva, including a raising of the head and/or tail of the animal or a full-body contraction toward the middle of the body. When stimuli are applied to the ventral body surface, at and below 14°C, a full-body contraction is the predominant response. CIII md neurons are activated by noxious cold stimuli, and optogenetic activation of these neurons is sufficient to induce cold-evoked contraction behavior (Turner et al., 2016). Like CIV neurons, the dendritic arbors of CIII neurons tile the epidermal wall, and the axonal projections of CIII neurons terminate in the VNC, immediately lateral to those of CIV neurons (Grueber et al., 2002, 2007).
Drosophila larvae also display nociceptive sensitization to noxious thermal stimuli. Both thermal allodynia and hyperalgesia can be induced in larvae by application of ultraviolet (UV) radiation, which results in epidermal damage (Babcock, Landry, & Galko, 2009). Following UV radiation, larvae roll in response to temperatures as low as 34°C—the severity of allodynia peaks at 24 hours after radiation and vanishes within 48 hours (Babcock et al., 2009). Larvae also respond faster to a 45°C stimulus—the severity of hyperalgesia peaks at 8 hours after radiation and vanishes within 24 hours (Babcock et al., 2009).
Adult Drosophila are also able to detect noxious heat; withdrawal responses are generally measured by monitoring thermal preference/avoidance in an experimental chamber having both noxious and nonnoxious temperature zones (Neely et al., 2010; Aldrich, Kasuya, Faron, Ishimoto, & Kitamoto, 2010), as well as by jump latency, when presented with a noxious heat stimulus via laser or hot plate assay (Xu et al., 2006). The cellular basis of heat nociception in adults remains largely unknown. C. elegans is robust with regard to what thermal environments it can inhabit; worms are able to optimally reproduce in temperatures up to 20°C, and they are able to adapt to new stable temperatures in as little as a few hours (Harvey & Viney, 2007; Hedgecock & Russell, 1975). Despite the robust nature of C. elegans, thermoregulation is important to maximizing fitness—like other ectotherms, C. elegans maintains a stable internal temperature by modulating its behavior (Garrity, Goodman, Samuel, & Sengupta, 2010). Temperature-dependent behaviors in C. elegans were first described in Hedgecock and Russell (1975), who observed that a worm will tend to move toward the temperature at which it was reared, thereafter seeking to stay at that temperature.
The neuroethology of C. elegans thermoregulation has been documented in great detail (Clark, Gabel, Lee, & Samuel, 2007; Colosimo et al., 2004; Garrity et al., 2010; Glauser, 2013; Jurado, Kodama, Tanizawa, & Mori, 2010; Luo, Clark, Biron, Mahadevan, & Samuel, 2006; Ramot, MacInnis, Lee, & Goodman, 2008; Ryu & Samuel, 2002). C. elegans responds to noxious heat in a body-location-specific manner. A head stimulus initiates a three-part withdrawal response consisting of backward locomotion, a so-called omega turn, and subsequent forward locomotion (Wittenburg & Baumeister, 1999). The omega turn is characterized by the head of the animal curling back and crossing or touching the tail—in order to meet the criteria for such a turn, with the animal’s body divided into thirds, the angle of the head segment should exceed 30° (Yemini, Jucikas, Grundy, Brown, & Schafer, 2013). Response to tail stimulation is dependent on the locomotive state of the animal: when the animal is stationary, it will initiate backward locomotion, while a forward-moving animal will accelerate in its movement (Wittenburg & Baumeister, 1999). While head and tail stimulus-responses are demonstrably deterministic, studies have described a probabilistic response to midbody stimulation, consisting of either forward locomotion, backward locomotion, or pausing (Mohammadi, Byrne Rodgers, Kotera, & Ryu, 2013).
Traditionally, and mimicking mammalian literature (Basbaum, Bautista, Scherrer, & Julius, 2009), temperatures in excess of 30°C have been used to elicit nocifensive behaviors (Chatzigeorgiou et al., 2010; Glauser et al., 2011; Liu et al., 2012; Wittenburg & Baumeister, 1999). However, recent work has demonstrated that these behaviors can be elicited via very small changes in absolute temperature (≤1.4°C); the response has been suggested to be modulated, in a dose-dependent fashion, by the rate of change of temperature across the body of the animal, rather than by the crossing of a temperature threshold (Mohammadi et al., 2013). Noxious cold has been rarely studied in C. elegans; however, exposure to acute cold shock (shift from 20°C to 15°C) results in an increased incidence of omega turns (Chatzigeorgiou et al., 2010).
Liu et al. (2012) have implicated several specific neurons as primary noxious heat nociceptors: In the head, AFD and FLP neurons, and in the tail, PHC neurons. AFD neurons are single-rod-shaped, ciliated neurons found in the amphids, a pair of chemosensory organs located in the head, whereas FLP neurons are multidendritic sensory neurons that cover the head and neck of the animal (Hall & Treinin, 2011). PHC neurons are sexually dimorphic neurons found in the tail spike of the worm (Sulston, Albertson, & Thomson, 1980). AIB interneurons, a pair of first-layer amphid interneurons, have been demonstrated to be important to thermal nociception, and are electrically coupled to AFD (Liu, Schulze, & Baumeister, 2012; White et al., 1986). AIB interneurons are thought to be a site of sensory integration, and the pair synapse directly onto motor neurons (Wakabayashi, Kitagawa, & Shingai, 2004). In addition, RMG interneurons, which synapse onto neuromuscular junctions in the head, play a role in sensitivity to noxious heat (Glauser et al., 2011). Concerning noxious cold detection, Chatzigeorgiou et al. (2010) have demonstrated the necessity of PVD neurons.
Molecular Regulation of Thermal Nociception and Sensitization
Two TRP family channels are known to be involved in heat nociception: Painless and dTRPA1. These TRPs are multimodal, as they also detect noxious mechanical stimuli. Painless acts in the detection of noxious heat via larval CIV neurons (Tracey Jr. et al., 2003; Hwang et al., 2012), and it is likewise required in adult thermal nociception in painless-expressing neurons outside the mushroom bodies (Neely et al., 2011; Xu et al., 2006). Interestingly, expression of canonical Painless, which bears N-terminal ARs—typically associated with mechanical gating—appears necessary for the proper function of Painless in thermal, but not mechanical, nociception (Hwang et al., 2012). dTRPA1 is required in both adult and larval thermal nociception (Babcock et al., 2011; Neely et al., 2011; Zhong et al., 2012). dTRPA1 has been implicated in sensing rapid changes in temperature; such changes initiate rolling at lower-than-typical threshold temperatures (Luo, Shen, & Montell, 2017). Knockdowns of dTRPA1 result in severe disruptions to an animal’s ability to detect and respond to noxious thermal stimuli. At least four isoforms of dTRPA1 exist, and surprisingly, one isoform, dTRPA1-C, is not a primary heat sensor, yet it is necessary and sufficient for rescuing thermal nociception defects; expression of the thermosensory isoform, dTRPA1-A, in CIV neurons leads to thermal allodynia and appears dependent on the presence of a heat-sensing domain referred to as a TRP Ankyrin Cap (TAC) element (Zhong et al., 2012).
Many other genes have been implicated in heat nociception. One of them, subdued, a member of the anoctamin family of calcium-activated chloride channels, is required in both adults and larvae; mutants exhibits nociceptive defects on the same order of magnitude as dTRPA1 and painless mutants (Jang et al., 2015). The amnesiac gene is predicted to code a precursor to a neuropeptide and has been associated with a variety of functions, including thermal preference (Hong et al., 2008). The amnesiac gene is required for thermal nociception in both adults and larvae, though it is not clear how it functions (Aldrich et al., 2010). The ecdysone steroid hormone receptor EcRA is also involved in modulating sensitivity to noxious thermal and mechanical stimuli, as CIV suppression of EcRA leads to hyposensitivity (McParland, Follansbee, Vesenka, Panaitiu, & Ganter, 2015). A genomewide, pan-neural screen for mediators of thermal nociception in adults identified hundreds of genes with aberrant responses to noxious heat, including straightjacket (stj) gene, which encodes a member of the α2δ family of voltage-gated Ca2+ channels (Neely et al., 2010). The stj gene is expressed in both larvae and adults, and is necessary for generating a response to noxious heat. This nociceptive function is evolutionarily conserved as mice mutant for the stj ortholog CACNA2D3 (α2δ3) are behaviorally defective for heat pain sensitivity, and humans with SNP variants in α2δ3 exhibit reduced sensitivity to acute noxious heat and chronic back pain (Neely et al., 2010). In addition, a screen of CIV nociceptor enriched genes identified a host of previously uncharacterized and evolutionarily conserved genes that, when disrupted via CIV-specific RNAi, are either noxious heat insensitive or hypersensitive, and in many cases, they disrupt CIV dendritic morphology. Nociceptive insensitivity was often associated with reductions in dendritic field coverage, whereas hypersensitivity was, in some cases, associated with enhanced coverage (Honjo et al., 2016). Moreover, genomewide RNAi screens in adult flies have identified additional genes involved in heat nociception—40 of these genes have been identified as having orthologous function in pain (Neely et al., 2012).
The characteristic bursting pattern observed during thermal activation of CIV neurons requires the proper expression of both dTRPA1 and Painless (Terada et al., 2016; Xiang, Yuan, Vogt, Looger, Jan, & Jan, 2010). It appears that these two channels operate in a coordinated but nonadditive fashion (Terada et al., 2016; Xiang et al., 2010). The Ca2+ transient intrinsically linked to this bursting activity requires the proper expression of the L-type voltage-gated Ca2+ channel (VGCC). Following the application of noxious heat, L-type VGCC-dependent spikes drive the inward Ca2+ transient (Terada et al., 2016). This transient is a necessary component of thermal nociception, but it is not required for general CIV excitability, such as when detecting aversive light (Terada et al., 2016; Xiang et al., 2010).
Drosophila cold nociception requires at least three TRP channels: Pkd2, NompC, and Trpm (Figure 1). Each of these TRP channels function at the sensory transduction stage in mediating cold-evoked contraction behavior. Moreover, Pkd2 is able to confer cold sensitivity to nonsensitive cells and therefore is thought to be a direct cold sensor. CIII neurons exhibit increases in intracellular Ca2+ concentrations following noxious cold stimulation, and disruptions in the function of these TRP channels variably affects these cold-induced Ca2+ responses. Cold-induced Ca2+ responses are decreased in Pkd2 mutants, whereas Trpm and nompC mutants exhibited variable increases in Ca2+ responses to cold, suggesting potentially complex roles for NompC and Trpm in regulating CIII calcium homeostasis upon cold exposure (Turner et al., 2016). This study also implicates NompC as a multimodal TRP channel mediating both gentle touch and noxious cold behaviors; however, it is not yet known if the ARs that mediate gating in response to gentle touch are required for NompC-mediated gating in response to cold (Turner et al., 2016).
UV-induced thermal nociceptive sensitization is thought to occur via multiple signaling mechanisms that converge on Painless/dTRPA1. UV-induced damage to epidermal cells results in the activation of the Dronc caspase and the production of the tumor necrosis factor (TNF) homolog, Eiger, which is released from epidermal cells. Eiger then binds to the TNF Receptor homolog, Wengen, on CIV nociceptors, to elicit thermal allodynia (Babcock et al., 2009). In parallel to Eiger/Wengen signaling, Hedgehog (Hh) signaling likewise regulates nociceptive sensitization and is required for both thermal allodynia and hyperalgesia following UV-induced tissue damage. Hh signaling acts through Painless for thermal allodynia, and via dTRPA1 for thermal hyperalgesia; Hh analgesic signaling appears conserved in mammals (Babcock et al., 2011). Neuropeptide signaling has also been implicated in UV-induced thermal allodynia, whereby UV damage leads to the release of the Substance P–related neuropeptide, Tachykinin (DTK) (Im et al., 2015). The working model for DTK function upstream of Hh signaling involves the release of DTK from brain neurons that target CIV nociceptors expressing the DTK receptor (DTKR), which is coupled to trimeric G proteins that initiate a signaling cascade. This cascade facilitates CIV release of Hh, via Dispatched. Hh acts in an autocrine manner on CIV neurons to bind the Patched receptor, relieving Smoothened inhibition and activating, or modulating, Painless, to promote thermal allodynia (Im et al., 2015).
In C. elegans, at least three channels play some role in thermal nociception: the downstream cyclic nucleotide-gated (CNG) channel of the tax-2 and tax-4 genes, and two TRPV channels, OCR-2 and OSM-9 (Liu et al., 2012). In AFD neurons, tax-2 and tax-4 are necessary, whereas in FLP and PHC neurons, TRPV channel expression is necessary (Liu et al., 2012). There is no evidence that the tax-2/tax-4 CNG channel is expressed in the tail, and yet disruptions in these genes cause defects in tail stimulus–associated responses. The RMG interneuron has been suggested to act in a TRPV-independent manner, via the G-protein coupled neuropeptide receptor NPR-1 (Glauser et al., 2011). Loss of NPR-1 or its receptor ligand FLP-21 results in an increased threshold by which escape behaviors are elicited in response to noxious heat (Glauser et al., 2011). Detection of acute cold in PVD neurons is thought to act via TRPA-1 and nAChR subunits. TRPA-1 expression can confer cold sensitivity to other cell types, suggesting that it is a primary sensory transducer for cold (Chatzigeorgiou et al., 2010), whereas mutants in the nAChR subunits DEG-3 and DES-2 impairs cold-evoked escape locomotion (Cohen et al., 2014).
Chemical Nociception and Aversion
As with other noxious stimuli, noxious chemicals can harm an animal quickly, irreparably, or both; this fact has facilitated the evolution of systems for detecting and responding to these dangers in a manner more immediate than passive aversion. A clear and widely accepted distinction between aversive chemicals and noxious chemicals has not yet been established in Drosophila literature. That said, some proteins known to contribute to mechanical and thermal nociception have also been implicated in sensing potentially noxious chemicals.
Two methods have been used to study chemical nociception and aversive chemosensation in adult Drosophila: the two-choice preference assay and a measure of the proboscis extension response (PER). In a two-choice preference assay, animals are given two food choices and their preferences are recorded—adult flies tend to avoid consuming foods that contain chemicals suspected to be noxious (Al-Anzi, Tracey Jr., & Benzer, 2006). The PER is characterized by the extension of the proboscis—the tubular feeding structure attached to the head—following leg contact with sugary solutions. Animals display a variety of disruptions to PER in response to suspected noxious chemicals, including a failure to fully extend, a quick retraction after contact, or inhibition of PER in trials subsequent to initial ingestion (Al-Anzi et al., 2006; K. Kang et al., 2010).
Many chemicals have been shown to be aversive via two-choice assays: papaverine, berberine, denatonium, quinine, caffeine, and strychnine, all bitter-tasting to humans; allyl- and benzyl-isothiocyanate (AITC and BITC), which contribute to the pungent taste of wasabi; and DEET, L-canacanine, citronella, lobeline, and camphor, a group of pesticides or suspected pest deterrents (Al-Anzi et al., 2006; Jeong et al., 2013; Lee et al., 2012; Lee, Kim, & Montell, 2010; Shim et al., 2015; Zhang, Ni, & Montell, 2013). Adult flies are also known to avoid high concentrations of salt (Alves, Sallé, Chaudy, Dupas, & Manière, 2014; Zhang et al., 2013). Fewer chemicals have been implicated in disruptions to the PER: AITC and BITC, N-methyl maleimide (NMM), and Cinnamaldehyde (CA) (Al-Anzi et al., 2006; K. Kang et al., 2010). These compounds are all reactive electrophiles, which are aversive to many animals and might damage a variety of biological molecules (Basbaum, Bautista, Scherrer, & Julius, 2009; K. Kang et al., 2010).
These studies, in Drosophila, are not without controversy. Im & Galko (2012) wrote the following:
There is ambiguity in [two-choice and PER] assays, however, as to whether they can distinguish a painful experience from an unpleasant taste. The adverse sensory experience of humans with these same compounds suggests that flies may perceive them as genuinely noxious although data on whether they can directly cause tissue damage in flies is needed.
In Drosophila, gustatory neurons found in the proboscis, internal pharyngeal sensory structures, leg tarsus, and wing have been suggested to be chemical nociceptors. These neurons are identified by the expression of Painless, and typically, heterogeneous expression of one of three gustatory receptors: Gr66a, Gr47a, or Gr32a (Al-Anzi et al., 2006). Neurons of the labral sense organ (LSO), a gustatory organ open to the esophageal lumen, have also been suggested to be nociceptors—these cells are identified by expression of the dTRPA1 channel (K. Kang et al., 2010). Gr66a neurons are broadly tuned avoidance detectors and have been specifically implicated in the detection of bitter compounds (Marella et al., 2006). These cells project to the medial subesophageal ganglion (SOG), a region of the ventral brain where all other gustatory neurons—excepting those of the ovipositor, wing, and parts of the leg—project (Power, 1948; Rajashekhar & Singh, 1994; Stocker & Schorderet, 1981). Depending on their location, Gr47a and Gr32a neurons project to either the SOG or to the thoracic ganglia (if found in the wings or legs) (Marella et al., 2006). Gr47a cells are known to detect strychnine, while Gr32a cells are involved in a variety of processes, including gustation, aggression, and courtship (Andrews et al., 2014; Lee, Moon, Wang, & Montell, 2015).
Like all animals, C. elegans has to navigate a complex environment and has developed robust chemosensory systems by which it can find food or avoid danger. In fact, more than 5% of C. elegans genes are thought to play a role in chemosensation (Bargmann, 2006).
Culotti and Russell (1978) were the first to describe how nematodes respond to noxious chemical stimuli: animals reverse-locomote, turn, and then change their navigational trajectory. C. elegans respond to a variety of different chemical stimuli that could be considered noxious: high osmolality, of at least both sugars and salts (Culotti & Russell, 1978); alkaloids, alkaloid derivatives, and at least one precursor to plant alkaloids, many of which are known to taste bitter to humans (Hilliard, Bergamasco, Arbucci, Plasterk, & Bazzicalupo, 2004); metals, such as Cd2+ and Cu2+ (Sambongi et al., 1999); highly acidic and basic solutions (Wang et al., 2016); detergents, such as sodium dodecyl sulfate (SDS; Hilliard, Bargmann, & Bazzicalupo, 2002); and a number of organic odorants (Chao, Komatsu, Fukuto, Dionne, & Hart, 2004).
Plasticity has been observed in how worms react to these various stimuli. Sensory adaptation—a decrease in sensitivity following repeated application of a stimulus—has been observed in worms (Hilliard et al., 2005). During periods of starvation, worms show increased adaptation, as well as decreased sensitivity to noxious stimuli, presumably such that a worm can have an increased area by which to find food in nutrient-poor environments. Conversely, during food-rich periods, worms display decreased adaptation and are more sensitive to these stimuli (Ezcurra, Walker, Beets, Swoboda, & Schafer, 2016).
In nematodes, chemosensation, including chemical nociception, is primarily mediated by ASH, a multimodal mechanosensory and chemosensory neuron found in the amphid (Bargmann, Thomas, & Horvitz, 1990; Kaplan & Horvitz, 1993; Hilliard et al., 2005). ASH neurons synapse with AVA, AVB, and AVD neurons, three of four pairs of command interneurons that synapse on ventral cord motor neurons (Ward, 1973). This network appears to function via activation of excitatory AMPA-type and NMDA-type glutamate receptors expressed in AVA, AVB, and AVD (Hart, Sims, & Kaplan, 1995; Maricq, Peckol, Driscoll, & Bargmann, 1995; Mellem, Brockie, Zheng, Madsen, & Maricq, 2002).
ASK and ADL are used in the detection of quinine (Hilliard et al., 2004); ASK and ASE are both activated by noxious heavy metals and detergents (Hilliard et al., 2002; Sambongi et al., 1999); finally, AWB (which is known to normally be involved in detecting aversive, but not noxious, odorants) and ADL are both recruited to perform nociceptive functions during periods of starvation (Chao et al., 2004; Troemel, Kimmel, & Bargmann, 1997). PHA and PHB are two pairs of ciliated neurons found in the phasmids, a pair of posterior sensory organs that synapse to the AVA, AVD, and PVC command interneurons, which collectively coordinate forward and backward locomotion (Chalfie et al., 1985; Maricq et al., 1995). PHA and PHB are both mechanosensory and chemosensory, and act as a means to, when necessary, inhibit backward locomotion initiated by amphid sensory neurons (Hilliard et al., 2002).
Sensory sensitization and adaptation, at least in response to Cu2+, are thought to be modulated by reciprocal inhibition between ASH and ASI, an amphid neuron with chemotactic functions (Guo et al., 2015). RIC interneurons and ADFs, a pair of sensory neurons with chemotactic functions, both appear to act as interneurons in this modulatory network (Guo et al., 2015). Serotonin is again important; SER-5 and SER-3 are expressed in ASH and ASI neurons, respectively, and act as the target and means of inhibition (Guo et al., 2015). During food-rich periods, worms show enhanced sensitivity to noxious chemicals and decreased levels of adaptation; this is thought to be a dopaminergic process acting through the AUA interneuron (Ezcurra, Tanizawa, Swoboda, & Schafer, 2011; Ezcurra et al., 2016).
Molecular Regulation of Chemical Nociception and Aversion
In flies, both Painless and dTRPA1 are implicated in chemical nociception (Al-Anzi et al., 2006; K. Kang et al., 2010), although the precise function that Painless plays in this process is unclear. Al-Anzi et al. (2006) reported that Painless is necessary for detecting AITC and BITC, whereas Kang et al. (2010) reported that painless mutants continue to detect chemicals such as AITC and NMM, but with decreased ability, suggesting Painless has some auxiliary or redundant function. Im and Galko (2012) suggested that these differences may be accounted for by differences in behavioral scoring protocols. The dTRPA1 is necessary for the detection of the noxious reactive electrophiles AITC, NMM, and CA (K. Kang et al., 2010; Zhong et al., 2012). The dTRPA1-B/C isoforms displayed more specific activation responses to AITC than noxious heat (Zhong et al., 2012). The dTRPA1 also appears necessary for detecting citronella and aristolochic acid, but it does not appear to function as a direct sensor (Kim et al., 2010; Kwon et al., 2010).
Three molecules have been implicated in the detection of aversive salt concentrations (≥200mM): the ionotropic receptor IR76b, which is found in many sensory sensilla; and Serrano and PPK19, both implicated in the activity displayed by gustatory neurons following salt stimulation (Alves et al., 2014; Zhang et al., 2013). A number of chemical stimuli, which are at the very least aversive, are detected by gustatory receptors (indicated by gene name): L-canavarine, by GR8a, GR66a, and GR98b; strychnine, by GR47a, GR32a, GR33a, and GR66a; caffeine, by GR66a, GR93a, and GR33a; DEET, by GR32a, GR33a, and GR66a; papavarine, by GR32a, GR33a, GR66a; and lobeline, by GR32a, GR33a, and GR66a (Lee, Moon, & Montell, 2009; Lee et al., 2010, 2012, 2015; Moon, Köttgen, Jiao, Xu, & Montell, 2006; Shim et al., 2015). The odorant binding protein OBP49a is expressed in thecogen cells—nonneuronal accessory cells found in the sheath of L- and S-type sensilla—where it functions to modulate the activity of gustaory neurons by suppressing feeding activity when bitter compounds are detected alongside sucrose (Jeong et al., 2013). Odorant receptors have also been implicated in avoiding aversive chemicals (Li & Liberles, 2015).
Circuits involved in chemical nociception have been more thoroughly characterized in C. elegans. Following exposure of ASH to noxious chemical stimuli, intracellular Ca2+ concentrations are dramatically increased. At least three genes have been implicated as necessary, in most cases, to generate this Ca2+ transient: osm-9, which encodes a TRPV-type channel; egl-19, which encodes a pore-forming subunit of a voltage-gated calcium channel; and odr-3, which encodes 1 of 20 G-protein alpha-subunits and is required for a variety of functions, including cilia morphogenesis (Hilliard et al., 2005). Ca2+ release from intracellular stores, via IP3R and RYR, has also been suggested as a component of this transient (Baker, Edwards, & Rickard, 2013; Zahratka, Williams, Summers, Komuniecki, & Bamber, 2015). Given that at least a fraction of Ca2+ enters ASH through voltage gated Ca2+ channels, and that proper cilia formation is required for nociceptive function (at least in the case of heavy metals), it has been suggested that noxious chemicals directly depolarize ASH (Hilliard et al., 2004, 2005). Some recent evidence, however, has indicated that Ca2+ transient amplitudes may not be reliably predictive of cellular activity—inhibition of voltage-gated Ca2+ channels did not inhibit nocifensive behaviors in animals challenged with 1-octanol, a noxious odorant (Baker et al., 2013; Zahratka et al., 2015).
Many other genes have been associated with noxious chemical detection (many with functions specific to particular stimuli). As with thermal nociception, the proper expression of tax-2 and tax-4 genes appears necessary for normal nonnoxious chemical sensation (Colbert, Smith, & Bargmann, 1997). In addition, tax-4 mutants show a decreased ability to detect SDS, a noxious detergent (Hilliard et al., 2002). The gpa-3 gene, which encodes another G-protein alpha-subunit, and gpc-1, which encodes one of two G-protein γ-subunits, both appear to be involved in quinine detection; meanwhile, gpa-13, gpa-15, and odr-3 appear largely dispensable in this process (Hilliard et al., 2004). The qui-1 code encodes a protein involved in sensing alkaloids (quinine), detergents (SDS), and acidic pH that is expressed in ASH and ADL neurons (Hilliard et al., 2004). The tmc-1 gene encodes a transmembrane channellike protein thought to be an Na+-sensitive channel; TMC-1 works in conjunction with OSM-9 to specifically facilitate the detection of alkaline environments (Wang, Li, Liu, Liu, & Xu, 2016).
Sensory adaptation—the reduction in sensitivity to a particular stimulus given persistent presentation—has been observed in ASH neurons following application of Cu2+ (Hilliard et al., 2005). The gpc-1 gene, which is used in quinine nociception and is required for chemoattractant adaptation, has been found to be necessary for nociceptive adaptation (Hilliard et al., 2005; Jansen, Weinkove, & Plasterk, 2002). Decreases in sensitivity and increases in adaptation during periods of starvation are thought to be controlled by NPR-1 and NPR-2, two members of the neuropeptide F/Y family. Dopaminergic increases of sensitivity and decreases in adaptation—seen during food-rich periods—are NPR-1-dependent processes (Ezcurra et al., 2016).
Multimodality and Nociception
Invertebrate nociceptive systems are not always instantiated in stimulus-unique neural circuitry. In fact, nociceptors and their downstream circuits are often multimodal—they transduce, process, and generate relevant behavioral responses to a variety of sensory information. Understanding how these systems function may yield some generalizable principles by which to understand a number of multimodal sensory systems.
In Drosophila larvae, CIII and CIV md neurons are both multimodal. CIV neurons function as multimodal thermal and mechanical nociceptors, and as phototransducers, they enable larva to sense light over the full body and to exhibit photoavoidance of potentially dangerous light stimuli (e.g., UV irradiation) (Xiang et al., 2010). CIII neurons respond to both innocuous mechanical stimulation (Tsubouchi et al., 2012; Yan et al., 2013) and noxious cold (Turner et al., 2016).
CIV multimodality can be explained, at least in part, by the way that they electrically respond to various stimuli (Terada et al., 2016). Noxious heat induces high-frequency bursting interrupted by pauses of increasing length, correlated with an increase in intracellular dendritic Ca2+. This response pattern is unique to thermal stimulation and contributes to the distinct behaviors observed in response to noxious heat (rolling) versus light avoidance (Terada et al., 2016; Xiang et al., 2010).
In the case of CIII multimodality, behavioral output is dependent upon activation levels. Low levels of CIII activation predominantly initiate gentle-touch responses, while high levels predominantly initiate noxious cold responses. These findings are corroborated by calcium imaging, suggesting that CIII neurons function as low-threshold mechanosensors and high-threshold nociceptors (Turner et al., 2016). Intriguingly, with respect to the organization of thermosensory nociceptors, coactivation studies of CIII and CIV neurons reveal predominant cold-evoked behavior, suggesting that noxious cold signals and behavior override those mediated by CIV heat nociceptors when both neurons are simultaneously activated (Turner et al., 2016).
Drosophila sensory neurons converge on complex multimodal circuitry, and information often converges at several layers, making complex behavioral selection possible. For example, CIV md neurons and vibration-sensitive mechanosensory neurons both share targets with, and independently converge on, a set of interneurons called Basins. Basins transmit sensory information through independent, multilayered nerve-cord and brain pathways, and information ultimately converges on command like Goro neurons in the motor nerve cord (Ohyama et al., 2015). Nociceptive systems also converge with one another; when costimulating CIII and CIV neurons, the CIII-specific contraction response predominantly occurs (Turner et al., 2016). The mechanism by which this behavior is selected has not yet been elucidated. Many circuit motifs—including reciprocal inhibition of inhibition, lateral disinhibition, and feedback disinhibition—have been implicated in behavior selection following innocuous stimuli (Jovanic et al., 2016); however, it is not yet clear how or if these circuit motifs are involved in processing noxious stimuli.
A large number of C. elegans sensory and interneurons are multimodal, and some respond to a wide range of both noxious and innocuous stimuli. For example, ASH neurons respond to gentle mechanical stimuli, hyperosmolarity, volatile chemicals, heavy metals, detergents, alkaline pH, and alkaloids (Bargmann et al., 1990; Kaplan & Horvitz, 1993; Hilliard et al., 2005).
ASH neurons respond to differing stimuli via a variety of signaling mechanisms: tmc-1 is required for detecting alkaline pH and salt sensation (Chatzigeorgiou et al., 2013); glr-1, an ionotropic glutamate receptor, is necessary for responding to nose-touch (Maricq et al., 1995); and itr-1, a putative inositol triphosphate receptor, is required for nose-touch and volatile chemical detection (Walker et al., 2009). Serotonin also may be important to ASH multimodality; sensitivity to octanol is decreased when serotonin levels are decreased (and increased when serotonin levels are increased), and following a gentle touch, Ca2+ transients can be generated only when serotonin is present (Chao et al., 2004; Hilliard et al., 2005). ASH multimodality also might be explained, in part, by high- or low-threshold activation; chemical stimuli activate ASH more strongly than mechanical stimuli (Hart et al., 1995; Maricq et al., 1995; Mellem et al., 2002).
Like Drosophila, C. elegans behavior selection circuits exhibit complex connectivity patterns. A multitude of sensory neurons converge on some combination of four pairs of interneurons, which themselves converge on ventral nerve cord motor neurons (the specifics of which are discussed in the “Mechanical Nociception,” “Thermal Nociception and Sensitization,” and “Chemical Nociception and Aversion” sections of this article). Circuits are also structured to drive behaviors in a stimulus-location-specific fashion—for example, chemosensory information from the head and tail are integrated into a head-to-tail spatial map of the chemical environment, which drives appropriate escape responses (Hilliard et al., 2002).
Many of the specific mechanisms of multimodal sensory processing remain to be elucidated. However, these examples demonstrate the complexity of the molecular, cellular, and circuit factors involved in such processing.
C. elegans and Drosophila have a multitude of sensory systems, many of which have been explored in great detail, including innocuous and noxious chemosensation; gentle and harsh mechanosensation; cool, warm, cold, and hot thermosensation; and photosensation. Despite this progress, many open questions exist regarding the specific mechanisms of, and interactions that exist between, nociceptive systems and their nonnoxious counterparts. Of particular interest is how these systems drive distinct and relevant behaviors in response to varying sensory inputs. Perhaps at a higher level, the extent that behavior selection can be modified in light of environmental cues remains to be described in detail. Answering these questions will not only serve to better our understanding of these individual species, but also may shed light on such systems in other animals, perhaps including humans.
Authors N.J.H. and A.A.P. contributed equally to this work. This work is supported in part by NINDS R01 NS086082, a Georgia State University (GSU) Brains and Behavior seed grant, and a 2CI Neurogenomics and Molecular Basis of Disease award (GSU) to D.N.C. A.A.P. was supported by a 2CI Neurogenomics Fellowship (GSU).
Al-Anzi, B., Tracey, W. D., Jr., & Benzer, S. (2006). Response of Drosophila to wasabi is mediated by painless, the fly homolog of mammalian TRPA1/ANKTM1. Current Biology, 16(10), 1034–1040.Find this resource:
Aldrich, B. T., Kasuya, J., Faron, M., Ishimoto, H., & Kitamoto, T. (2010). The amnesiac gene is involved in the regulation of thermal nociception in Drosophila melanogaster. Journal of Neurogenetics, 24(1), 33–41.Find this resource:
Alves, G., Sallé, J., Chaudy, S., Dupas, S., & Manière, G. (2014). High-NaCl perception in Drosophila melanogaster. The Journal of Neuroscience, 34(33), 10884–10891.Find this resource:
Andrews, J. C., Fernández, M. P., Yu, Q., Leary, G. P., Leung, A. K. W., Kavanaugh, M. P., et al. (2014). Octopamine neuromodulation regulates Gr32a-linked aggression and courtship pathways in Drosophila males. PLoS Genetics, 10(5), e1004356.Find this resource:
Árnadóttir, J., & Chalfie, M. (2010). Eukaryotic mechanosensitive channels. Annual Review of Biophysics, 39(1), 111–137.Find this resource:
Babcock, D. T., Landry, C., & Galko, M. J. (2009). Cytokine signaling mediates UV-induced nociceptive sensitization in Drosophila larvae. Current Biology, 19(10), 799–806.Find this resource:
Babcock, D. T., Shi, S., Jo, J., Shaw, M., Gutstein, H. B., & Galko, M. J. (2011). Hedgehog signaling regulates nociceptive sensitization. Current Biology, 21(18), 1525–1533.Find this resource:
Baker, K. D., Edwards, T. M., & Rickard, N. S. (2013). The role of intracellular calcium stores in synaptic plasticity and memory consolidation. Neuroscience & Biobehavioral Reviews, 37(7), 1211–1239.Find this resource:
Bargmann, C. I., Thomas, J. H., & Horvitz, H. R. (1990). Chemosensory cell function in the behavior and development of Caenorhabditis elegans. Cold Spring Harbor Symposia on Quantitative Biology, 55, 529–538.Find this resource:
Bargmann, C. I. (2006). Chemosensation in C. elegans, WormBook, ed. The C. elegans Research Community, WormBook.
Basbaum, A. I., Bautista, D. M., Scherrer, G., & Julius, D. (2009). Cellular and molecular mechanisms of pain. Cell, 139(2), 267–284.Find this resource:
Bechstedt, S., & Howard, J. (2008). Hearing mechanics: a fly in your ear. Current Biology, 18(18), R869–R870.Find this resource:
Bianchi, L., Gerstbrein, B., Frokjaer-Jensen, C., Royal, D. C., Mukherjee, G., Royal, M. A., et al. (2004). The neurotoxic MEC-4(d) DEG/ENaC sodium channel conducts calcium: Implications for necrosis initiation. Nature Neuroscience, 7(12), 1337–1344.Find this resource:
Bounoutas, A., & Chalfie, M. (2007). Touch sensitivity in Caenorhabditis elegans. Pflügers Archiv - European Journal of Physiology, 454(5), 691–702.Find this resource:
Carton, Y., & Frey, F. (1984). Analyse expérimentale de trois niveaux d’interactions entre Drosophila melanogaster et le parasite Leptopilina boulardi (sympatrie, allopatrie, xénopatrie). Génétique, Sélection, Évolution, 16(4), 417–430.Find this resource:
Cermenati, G., Giatti, S., Cavaletti, G., Bianchi, R., Maschi, O., Pesaresi, M., et al. (2010). Activation of the liver X receptor increases neuroactive steroid levels and protects from diabetes-induced peripheral neuropathy. Journal of Neuroscience, 30(36), 11896–901.Find this resource:
Chalfie, M., Sulston, J., White, J., Southgate, E., Thomson, J., & Brenner, S. (1985). The neural circuit for touch sensitivity in Caenorhabditis elegans. Journal of Neuroscience, 5(4), 956–964.Find this resource:
Chao, M. Y., Komatsu, H., Fukuto, H. S., Dionne, H. M., & Hart, A. C. (2004). Feeding status and serotonin rapidly and reversibly modulate a Caenorhabditis elegans chemosensory circuit. Proceedings of the National Academy of Sciences USA, 101(43), 15512–15517.Find this resource:
Chatzigeorgiou, M., Bang, S., Hwang, S. W., & Schafer, W. R. (2013). Tmc-1 encodes a sodium-sensitive channel required for salt chemosensation in C. elegans. Nature, 494, 95–99.Find this resource:
Chatzigeorgiou, M., Yoo, S., Watson, J. D., Lee, W.-H., Spencer, W. C., Kindt, K. S., et al. (2010). Specific roles for DEG/ENaC and TRP channels in touch and thermosensation in C. elegans nociceptors. Nature Neuroscience, 13(7), 861–868.Find this resource:
Cho, H., & Oh, U. (2013). Anoctamin 1 mediates thermal pain as a heat sensor. Current Neuropharmacology, 11(6), 641–651.Find this resource:
Clark, D. A., Gabel, C. V., Lee, T. M., & Samuel, A. D. T. (2007). Short-term adaptation and temporal processing in the cryophilic response of Caenorhabditis elegans. Journal of Neurophysiology, 97(3), 1903–1910.Find this resource:
Cohen, E., Chatzigeorgiou, M., Husson, S. J., Steuer-Costa, W., Gottschalk, A., Schafer, W. R., & Treinin, M. (2014). Caenorhabditis elegans nicotinic acetylcholine receptors are required for nociception. Molecular and Cellular Neurosciences, 59, 85–96.Find this resource:
Colbert, H. A., Smith, T. L., & Bargmann, C. I. (1997). OSM-9, a novel protein with structural similarity to channels, is required for olfaction, mechanosensation, and olfactory adaptation in Caenorhabditis elegans. Journal of Neuroscience, 17(21), 8259–8269.Find this resource:
Colosimo, M. E., Brown, A., Mukhopadhyay, S., Gabel, C., Lanjuin, A. E., Samuel, A. D. T., & Sengupta, P. (2004). Identification of thermosensory and olfactory neuron-specific genes via expression profiling of single neuron types. Current Biology, 14(24), 2245–2251.Find this resource:
Coste, B., Xiao, B., Santos, J.S., Syeda, R., Grandl, J., Spencer, K.S., et al. (2012). Piezo proteins are pore-forming subunits of mechanically activated channels. Nature, 483, 176–181.Find this resource:
Crook, R. J., & Walters, E. T. (2011). Nociceptive behavior and physiology of molluscs: Animal welfare implications. ILAR Journal, 52(2), 185–195.Find this resource:
Culotti, J. G., & Russell, R. L. (1978). Osmotic avoidance defective mutants of the nematode Caenorhabditis elegans. Genetics, 90(2), 243–256.Find this resource:
Ezcurra, M., Tanizawa, Y., Swoboda, P., & Schafer, W. R. (2011). Food sensitizes C. elegans avoidance behaviours through acute dopamine signaling. EMBO Journal, 30(6), 1110–1122.Find this resource:
Ezcurra, M., Walker, D. S., Beets, I., Swoboda, P., & Schafer, W. R. (2016). Neuropeptidergic signaling and active feeding state inhibit nociception in Caenorhabditis elegans. Journal of Neuroscience, 36(11), 3157–3169.Find this resource:
Garrity, P. A., Goodman, M. B., Samuel, A. D., & Sengupta, P. (2010). Running hot and cold: Behavioral strategies, neural circuits, and the molecular machinery for thermotaxis in C. elegans and Drosophila. Genes & Development, 24(21), 2365–2382.Find this resource:
Glauser, D. A. (2013). How and why Caenorhabditis elegans uses distinct escape and avoidance regimes to minimize exposure to noxious heat. Worm, 2(4), e27285.Find this resource:
Glauser, D. A., Chen, W. C., Agin, R., MacInnis, B. L., Hellman, A. B., Garrity, P. A., et al. (2011). Heat avoidance is regulated by transient receptor potential (TRP) channels and a neuropeptide signaling pathway in Caenorhabditis elegans. Genetics, 188(1), 91–103.Find this resource:
Goodman, M. B. (2006). Mechanosensation in C. elegans. In WormBook (Ed.), The C. elegans Research Community, WormBook.Find this resource:
Gorczyca, D. A., Younger, S., Meltzer, S., Kim, S. E., Cheng, L., Song, W., et al. (2014). Identification of Ppk26, a DEG/ENaC channel functioning with Ppk1 in a mutually dependent manner to guide locomotion behavior in Drosophila. Cell Reports, 9(4), 1446–1458.Find this resource:
Grueber, W. B., Jan, L. Y., & Jan, Y. N. (2002). Tiling of the Drosophila epidermis by multidendritic sensory neurons. Development, 129(12), 2867–2878.Find this resource:
Grueber, W. B., Ye, B., Moore, A. W., Jan, L. Y., & Jan, Y. N. (2003). Dendrites of distinct classes of Drosophila sensory neurons show different capacities for homotypic repulsion. Current Biology, 13(8), 618–626.Find this resource:
Grueber, W. B., Ye, B., Yang, C. H., Younger, S., Borden, K., Jan, L. Y., & Jan, Y. N. (2007). Projections of Drosophila multidendritic neurons in the central nervous system: links with peripheral dendrite morphology. Development, 134, 55–64.Find this resource:
Guo, M., Wu, T.-H., Song, Y.-X., Ge, M.-H., Su, C.-M., Niu, W.-P., et al. (2015). Reciprocal inhibition between sensory ASH and ASI neurons modulates nociception and avoidance in Caenorhabditis elegans. Nature Communications, 6, 5655.Find this resource:
Guo, Y., Wang, Y., Wang, Q., & Wang, Z. (2014). The role of PPK26 in Drosophila larval mechanical nociception. Cell Reports, 9(4), 1183–1190.Find this resource:
Hall, D. H., & Treinin, M. (2011). How does morphology relate to function in sensory arborsTrends in Neurosciences, 34(9), 443–451.Find this resource:
Hart, A. C., Sims, S., & Kaplan, J. M. (1995). Synaptic code for sensory modalities revealed by C. elegans GLR-1 glutamate receptor. Nature, 378(6552), 82–85.Find this resource:
Harvey, S. C., & Viney, M. E. (2007). Thermal variation reveals natural variation between isolates of Caenorhabditis elegans. Journal of Experimental Zoology Part B: Molecular and Developmental Evolution, 308B(4), 409–416.Find this resource:
Hedgecock, E. M., & Russell, R. L. (1975). Normal and mutant thermotaxis in the nematode Caenorhabditis elegans. Proceedings of the National Academy of Sciences USA, 72(10), 4061–4065.Find this resource:
Hilliard, M. A., Apicella, A. J., Kerr, R., Suzuki, H., Bazzicalupo, P., & Schafer, W. R. (2005). In vivo imaging of C. elegans ASH neurons: Cellular response and adaptation to chemical repellents. EMBO Journal, 24(1), 63–72.Find this resource:
Hilliard, M. A., Bargmann, C. I., & Bazzicalupo, P. (2002). C. elegans responds to chemical repellents by integrating sensory inputs from the head and the tail. Current Biology, 12(9), 730–734.Find this resource:
Hilliard, M. A., Bergamasco, C., Arbucci, S., Plasterk, R. H. A., & Bazzicalupo, P. (2004). Worms taste bitter: ASH neurons, QUI-1, GPA-3, and ODR-3 mediate quinine avoidance in Caenorhabditis elegans. EMBO Journal, 23(5), 1101–1111.Find this resource:
Hong, S.-T., Bang, S., Hyun, S., Kang, J., Jeong, K., Paik, D., et al. (2008). cAMP signalling in mushroom bodies modulates temperature preference behaviour in Drosophila. Nature, 454(7205), 771–775.Find this resource:
Honjo, K., Mauthner, S. E., Wang, Y., Skene, J. H. P., & Tracey, W. D., Jr. (2016). Nociceptor-enriched genes required for normal thermal nociception. Cell Reports, 16(2), 295–303.Find this resource:
Huang, M., & Chalfie, M. (1994). Gene interactions affecting mechanosensory transduction in Caenorhabditis elegans. Nature, 367(6462), 467–470.Find this resource:
Husson, S. J., Costa, W. S., Wabnig, S., Stirman, J. N., Watson, J. D., Spencer, W. C., et al. (2012). Optogenetic analysis of a nociceptor neuron and network reveals ion channels acting downstream of primary sensors. Current Biology, 22(9), 743–752.Find this resource:
Hwang, R. Y., Stearns, N. A., & Tracey, W. D. (2012). The ankyrin repeat domain of the TRPA protein Painless is important for thermal nociception but not mechanical nociception. PLoS ONE, 7(1), e30090.Find this resource:
Hwang, R. Y., Zhong, L., Xu, Y., Johnson, T., Zhang, F., Deisseroth, K., & Tracey, W. D. (2007). Nociceptive neurons protect Drosophila larvae from parasitoid wasps. Current Biology, 17(24), 2105–2116.Find this resource:
Im, S. H., & Galko, M. J. (2012). Pokes, sunburn, and hot sauce: Drosophila as an emerging model for the biology of nociception. Developmental Dynamics, 241(1), 16–26.Find this resource:
Im, S. H., Takle, K., Jo, J., Babcock, D. T., Ma, Z., Xiang, Y., & Galko, M. J. (2015). Tachykinin acts upstream of autocrine Hedgehog signaling during nociceptive sensitization in Drosophila. eLife, 4, e10735.Find this resource:
Ishikawa, Y., & Kamikouchi, A. (2016). Auditory system of fruit flies. Hearing Research, 338, 1–8.Find this resource:
Jang, W., Kim, J. Y., Cui, S., Jo, J., Lee, B.-C., Lee, Y., et al. (2015). The Anoctamin family channel Subdued mediates thermal nociception in Drosophila. Journal of Biological Chemistry, 290(4), 2521–2528.Find this resource:
Jansen, G., Weinkove, D., & Plasterk, R. H. A. (2002). The G-protein γ subunit gpc-1 of the nematode C. elegans is involved in taste adaptation. EMBO Journal, 21(5), 986–994.Find this resource:
Jeong, Y. T., Shim, J., Oh, S. R., Yoon, H. I., Kim, C. H., Moon, S. J., & Montell, C. (2013). An odorant binding protein required for suppression of sweet taste by bitter chemicals. Neuron, 79(4), 725–737. http://dx.doi.org/10.1016/j.neuron.2013.06.025.Find this resource:
Jovanic, T., Schneider-Mizell, C. M., Shao, M., Masson, J. B., Denisov, G., Fetter, R. D., et al. (2016). Competitive disinhibition mediates behavioral choice and sequences in Drosophila. Cell, 167(3), 858–870.Find this resource:
Jurado, P., Kodama, E., Tanizawa, Y., & Mori, I. (2010). Distinct thermal migration behaviors in response to different thermal gradients in Caenorhabditis elegans. Genes, Brain, and Behavior, 9(1), 120–127.Find this resource:
Kang, K., Pulver, S. R., Panzano, V. C., Chang, E. C., Griffith, L. C., Theobald, D. L., et al. (2010). Analysis of Drosophila TRPA1 reveals an ancient origin for human chemical nociception. Nature, 464(7288), 597–600.Find this resource:
Kang, L., Gao, J., Schafer, W. R., Xie, Z., & Xu, X. Z. S. (2010). C. elegans TRP family protein TRP-4 is a pore-forming subunit of a native mechanotransduction channel. Neuron, 67(3), 381–391.Find this resource:
Kaplan, J. M., & Horvitz, H. R. (1993). A dual mechanosensory and chemosensory neuron in Caenorhabditis elegans. Proceedings of the National Academy of Sciences USA, 90(6), 2227–2231.Find this resource:
Kernan, M., Cowan, D., & Zuker, C. (1994). Genetic dissection of mechanosensory transduction: Mechanoreception-defective mutations of Drosophila. Neuron, 12(6), 1195–1206.Find this resource:
Kim, S. E., Coste, B., Chadha, A., Cook, B., & Patapoutian, A. (2012). The role of Drosophila Piezo in mechanical nociception. Nature, 483(7388), 209–212.Find this resource:
Kim, S. H., Lee, Y., Akitake, B., Woodward, O. M., Guggino, W. B., & Montell, C. (2010). Drosophila TRPA1 channel mediates chemical avoidance in gustatory receptor neurons. Proceedings of the National Academy of Sciences USA, 107(18), 8440–8445.Find this resource:
Kwan, K. Y., Allchorne, A. J., Vollrath, M. A., Christensen, A. P., Zhang, D.-S., Woolf, C. J., et al. (2006). TRPA1 contributes to cold, mechanical, and chemical nociception but is not essential for hair-cell transduction. Neuron, 50(2), 277–289.Find this resource:
Kwon, Y., Kim, S. H., Ronderos, D. S., Lee, Y., Akitake, B., Woodward, O. M., et al. (2010). Drosophila TRPA1 channel is required to avoid the naturally occurring insect repellent citronellal. Current Biology, 20(18), 1672–1678.Find this resource:
Lee, Y., Kang, M. J., Shim, J., Cheong, C. U., Moon, S. J., & Montell, C. (2012). Gustatory receptors required for avoiding the insecticide l-canavanine. Journal of Neuroscience, 32(4), 1429–1435.Find this resource:
Lee, Y., Kim, S. H., & Montell, C. (2010). Avoiding DEET through insect gustatory receptors. Neuron, 67(4), 555–561.Find this resource:
Lee, Y., Moon, S. J., & Montell, C. (2009). Multiple gustatory receptors required for the caffeine response in Drosophila. Proceedings of the National Academy of Sciences USA, 106(11), 4495–4500.Find this resource:
Lee, Y., Moon, S. J., Wang, Y., & Montell, C. (2015). A Drosophila gustatory receptor required for strychnine sensation. Chemical Senses, 40(7), 525–533.Find this resource:
Li, Q., & Liberles, S. D. (2015). Aversion and attraction through olfaction. Current Biology, 25, R120–R129.Find this resource:
Li, W., Kang, L., Piggott, B. J., Feng, Z., & Xu, X. Z. (2011). The neural circuits and sensory channels mediating harsh touch sensation in C. elegans. Nature Communications, 2, 315Find this resource:
Liu, L., Yermolaieva, O., Johnson, W. A., Abboud, F. M., & Welsh, M. J. (2003). Identification and function of thermosensory neurons in Drosophila larvae. Nature Neuroscience, 6(3), 267–273.Find this resource:
Liu, S., Schulze, E., & Baumeister, R. (2012). Temperature- and touch-sensitive neurons couple CNG and TRPV channel activities to control heat avoidance in Caenorhabditis elegans. PLoS ONE, 7(3), e32360.Find this resource:
Luo, L., Clark, D. A., Biron, D., Mahadevan, L., & Samuel, A. D. T. (2006). Sensorimotor control during isothermal tracking in Caenorhabditis elegans. Journal of Experimental Biology, 209(23), 4652–4662.Find this resource:
Luo, J., Shen, W. L., & Montell, C. (2017). TRPA1 mediates sensation of the rate of temperature change in Drosophila larvae. Nature Neuroscience, 20, 34–41.Find this resource:
Mantyh, P. W. (1997). Inhibition of hyperalgesia by ablation of lamina i spinal neurons expressing the substance p receptor. Science 278, 275–279.Find this resource:
Marella, S., Fischler, W., Kong, P., Asgarian, S., Rueckert, E., & Scott, K. (2006). Imaging taste responses in the fly brain reveals a functional map of taste category and behavior. Neuron, 49(2), 285–295.Find this resource:
Maricq, A. V., Peckol, E., Driscoll, M., & Bargmann, C. I. (1995). Mechanosensory signalling in C. elegans mediated by the GLR-1 glutamate receptor. Nature, 378(6552), 78–81.Find this resource:
Mauthner, S. E., Hwang, R. Y., Lewis, A. H., Xiao, Q., Tsubouchi, A., Wang, Y., et al. (2014). Balboa (PPK-26) binds to Pickpocket in vivo and is required for mechanical nociception in Drosophila larvae. Current Biology, 24(24), 2920–2925.Find this resource:
McParland, A. L., Follansbee, T. L., Vesenka, G. D., Panaitiu, A. E., & Ganter, G. K. (2015). Steroid receptor isoform expression in Drosophila nociceptor neurons is required for normal dendritic arbor and sensitivity. PLoS ONE, 10(10), e0140785.Find this resource:
Mellem, J. E., Brockie, P. J., Zheng, Y., Madsen, D. M., & Maricq, A. V. (2002). Decoding of polymodal sensory stimuli by postsynaptic glutamate receptors in C. elegans. Neuron, 36(5), 933–944.Find this resource:
Mohammadi, A., Byrne Rodgers, J., Kotera, I., & Ryu, W. S. (2013). Behavioral response of Caenorhabditis elegans to localized thermal stimuli. BMC Neuroscience, 14, 66Find this resource:
Moon, S. J., Köttgen, M., Jiao, Y., Xu, H., & Montell, C. (2006). A taste receptor required for the caffeine response in vivo. Current Biology, 16(18), 1812–1817.Find this resource:
Moreau, N., Mauborgne, A., Bourgoin, S., Couraud, P.-O., Romero, I. A., Weksler, B. B., et al. (2016). Early alterations of Hedgehog signaling pathway in vascular endothelial cells after peripheral nerve injury elicit blood-nerve barrier disruption, nerve inflammation, and neuropathic pain development. Pain, 157(4), 827–39.Find this resource:
Neely, G. G., Hess, A., Costigan, M., Keene, A. C., Goulas, S., Langeslag, M., et al. (2010). A genome-wide Drosophila screen for heat nociception identifies α2δ3 as an evolutionary-conserved pain gene. Cell, 143(4), 628–638.Find this resource:
Neely, G. G., Keene, A. C., Duchek, P., Chang, E. C., Wang, Q.-P., Aksoy, Y. A., et al. (2011). TrpA1 regulates thermal nociception in Drosophila. PLoS ONE, 6(8), e24343.Find this resource:
Neely, G. G., Rao, S., Costigan, M., Mair, N., Racz, I., Milinkeviciute, G., et al. (2012). Construction of a global pain systems network highlights phospholipid signaling as a regulator of heat nociception. PLoS Genetics, 8(12), e1003071.Find this resource:
O’Hagan, R., Chalfie, M., & Goodman, M. B. (2005). The MEC-4 DEG/ENaC channel of Caenorhabditis elegans touch receptor neurons transduces mechanical signals. Nature Neuroscience, 8(1), 43–50.Find this resource:
Ohyama, T., Schneider-Mizell, C. M., Fetter, R. D., Aleman, J. V., Franconville, R., Rivera-Alba, M., et al. (2015). A multilevel multimodal circuit enhances action selection in Drosophila. Nature, 520(7549), 633–639.Find this resource:
Page, A. J., Brierley, S. M., Martin, C. M., Hughes, P. A., & Blackshaw, L. A. (2007). Acid sensing ion channels 2 and 3 are required for inhibition of visceral nociceptors by benzamil. Pain, 133(1–3), 150–60.Find this resource:
Panula, P., Aarnisalo, A. A., & Wasowicz, K. (1996). Neuropeptide FF, a mammalian neuropeptide with multiple functions. Progress in Neurobiology, 48(4), 461–487.Find this resource:
Power, M. E. (1948). The thoracico-abdominal nervous system of an adult insect, Drosophila melanogaster. Journal of Comparative Neurology, 88(3), 347–409.Find this resource:
Rajashekhar, K. P., & Singh, R. N. (1994). Neuroarchitecture of the tritocerebrum of Drosophila melanogaster. Journal of Comparative Neurology, 349(4), 633–645.Find this resource:
Ramot, D., MacInnis, B. L., Lee, H.-C., & Goodman, M. B. (2008). Thermotaxis is a robust mechanism for thermoregulation in C. elegans nematodes. Journal of Neuroscience, 28(47), 12546–12557.Find this resource:
Rosenzweig, M., Brennan, K. M., Tayler, T. D., Phelps, P. O., Patapoutian, A., & Garrity, P. A. (2005). The Drosophila ortholog of vertebrate TRPA1 regulates thermotaxis. Genes & Development, 19(4), 419–424.Find this resource:
Rosenzweig, M., Kang, K., & Garrity, P. A. (2008). Distinct TRP channels are required for warm and cool avoidance in Drosophila melanogaster. Proceedings of the National Academy of Sciences USA, 105(38), 14668–14673.Find this resource:
Ryu, W. S., & Samuel, A. D. T. (2002). Thermotaxis in Caenorhabditis elegans analyzed by measuring responses to defined thermal stimuli. Journal of Neuroscience, 22(13), 5727–5733.Find this resource:
Sambongi, Y., Nagae, T., Liu, Y., Yoshimizu, T., Takeda, K., Wada, Y., & Futai, M. (1999). Sensing of cadmium and copper ions by externally exposed ADL, ASE, and ASH neurons elicits avoidance response in Caenorhabditis elegans. NeuroReport, 10(4), 753–757.Find this resource:
Sandkühler, J. (2009). Models and mechanisms of hyperalgesia and allodynia. Physiological Reviews, 89(2), 707–758.Find this resource:
Sayeed, O., & Benzer, S. (1996). Behavioral genetics of thermosensation and hygrosensation in Drosophila. Proceedings of the National Academy of Sciences USA, 93(12), 6079–6084.Find this resource:
Sénatore, S., Rami Reddy, V., Sémériva, M., Perrin, L., & Lalevée, N. (2010). Response to mechanical stress is mediated by the TRPA channel Painless in the Drosophila heart. PLoS Genetics, 6(9), e1001088.Find this resource:
Seybold, V. S. (2009). The role of peptides in central sensitization. Handbook of Experimental Pharmacology, 194, 451–491.Find this resource:
Sherrington, C. S. (1906). The integrative action of the nervous system. New Haven, CT: Yale University Press.Find this resource:
Shim, J., Lee, Y., Jeong, Y. T., Kim, Y., Lee, M. G., Montell, C., & Moon, S. J. (2015). The full repertoire of Drosophila gustatory receptors for detecting an aversive compound. Nature Communications, 6, 8867.Find this resource:
Sneddon, L. U. (2015). Pain in aquatic animals. Journal of Experimental Biology, 218, 967–976.Find this resource:
Sorkin, L. S., Xial, W. H., Wagner, R. & Myers, R. R. (1997). Tumor necrosis factor-alpha induces ectopic activity in nociceptive primary afferent fibres. Neuroscience, 81, 255–262.Find this resource:
Staniland, A. A., & McMahon, S. B. (2009). Mice lacking acid-sensing ion channels (ASIC) 1 or 2, but not ASIC3, show increased pain behaviour in the formalin test. European Journal of Pain, 13(6), 554–63.Find this resource:
Stocker, R. F., & Schorderet, M. (1981). Cobalt filling of sensory projections from internal and external mouthparts in Drosophila. Cell and Tissue Research, 216(3), 513–523.Find this resource:
Sulkowski, M. J., Kurosawa, M. S., & Cox, D. N. (2011). Growing pains: Development of the larval nocifensive response in Drosophila. Biological Bulletin, 221(3), 300–306.Find this resource:
Sulston, J. E., Albertson, D. G., & Thomson, J. N. (1980). The Caenorhabditis elegans male: Postembryonic development of nongonadal structures. Developmental Biology, 78(2), 542–576.Find this resource:
Suzuki, M., Mizuno, A., Kodaira, K., & Imai, M. (2003). Impaired pressure sensation in mice lacking TRPV4. Journal of Biological Chemistry, 278(25), 22664–22668.Find this resource:
Terada, S.-I., Matsubara, D., Onodera, K., Matsuzaki, M., Uemura, T., & Usui, T. (2016). Neuronal processing of noxious thermal stimuli mediated by dendritic Ca(2+) influx in Drosophila somatosensory neurons. eLife, 5, e12959.Find this resource:
Tracey, W. D., Jr., Wilson, R. I., Laurent, G., & Benzer, S. (2003). Painless, a Drosophila gene essential for nociception. Cell, 113(2), 261–273.Find this resource:
Tsubouchi, A., Caldwell, J. C., & Tracey, W. D. (2012). Dendritic filopodia, Ripped Pocket, NOMPC, and NMDARs contribute to the sense of touch in Drosophila larvae. Current Biology, 22, 2124–2134.Find this resource:
Troemel, E. R., Kimmel, B. E., & Bargmann, C. I. (1997). Reprogramming chemotaxis responses: Sensory neurons define olfactory preferences in C. elegans. Cell, 91(2), 161–169.Find this resource:
Turner, H. N., Armengol, K., Patel, A. A., Himmel, N. J., Sullivan, L., Iyer, S. C., Bhattacharya, S., Iyer, S. P. R., et al. (2016). The TRP channels Pkd2, NompC, and Trpm act in cold-sensing neurons to mediate unique aversive behaviors to noxious cold in Drosophila. Current Biology, 26, 1–13.Find this resource:
Vogelstein, J. T., Park, Y., Ohyama, T., Kerr, R. A., Truman, J. W., Priebe, C. E., & Zlatic, M. (2014). Discovery of brainwide neural-behavioral maps via multiscale unsupervised structure learning. Science, 344, 386–392.Find this resource:
Wakabayashi, T., Kitagawa, I., & Shingai, R. (2004). Neurons regulating the duration of forward locomotion in Caenorhabditis elegans. Neuroscience Research, 50(1), 103–111.Find this resource:
Walker, D. S., Vázquez-Manrique, R. P., Gower, N. J. D., Gregory, E., Schafer, W. R., & Baylis, H. A. (2009). Inositol 1,4,5-trisphosphate signalling regulates the avoidance response to nose touch in Caenorhabditis elegans. PLoS Genetics, 5(9).Find this resource:
Wang, X., Li, G., Liu, J., Liu, J., & Xu, X. Z. S. (2016). TMC-1 mediates alkaline sensation in C. elegans through nociceptive neurons. Neuron, 91(1), 146–154.Find this resource:
Ward, S. (1973). Chemotaxis by the nematode Caenorhabditis elegans: Identification of attractants and analysis of the response by use of mutants. Proceedings of the National Academy of Sciences USA, 70(3), 817–821.Find this resource:
Way, J. C., & Chalfie, M. (1989). The mec-3 gene of Caenorhabditis elegans requires its own product for maintained expression and is expressed in three neuronal cell types. Genes & Development, 3(12a), 1823–1833.Find this resource:
Wieskopf, J. S., Mathur, J., Limapichat, W., Post, M. R., Al-Qazzaz, M., Sorge, R. E., et al. (2015). The nicotinic α6 subunit gene determines variability in chronic pain sensitivity via cross-inhibition of P2X2/3 receptors. Science Translational Medicine, 7(287), 287ra72.Find this resource:
White, J. G., Southgate, E., Thomson, J. N., & Brenner, S. (1986). The structure of the nervous system of the nematode Caenorhabditis elegans. Philosophical Transactions of the Royal Society of London B: Biological Sciences, 314(1165), 1–340.Find this resource:
Wicks, S. R., Roehrig, C. J., & Rankin, C. H. (1996). A dynamic network simulation of the nematode tap withdrawal circuit: Predictions concerning synaptic function using behavioral criteria. Journal of Neuroscience, 16(12), 4017–4031.Find this resource:
Wittenburg, N., & Baumeister, R. (1999). Thermal avoidance in Caenorhabditis elegans: An approach to the study of nociception. Proceedings of the National Academy of Sciences USA, 96(18), 10477–10482.Find this resource:
Xiang, Y., Yuan, Q., Vogt, N., Looger, L. L., Jan, L. Y., & Jan, Y. N. (2010). Light-avoidance-mediating photoreceptors tile the Drosophila larval body wall. Nature, 468(7326), 921–926.Find this resource:
Xu, S. Y., Cang, C. L., Liu, X. F., Peng, Y. Q., Ye, Y. Z., Zhao, Z. Q., & Guo, A. K. (2006). Thermal nociception in adult Drosophila: Behavioral characterization and the role of the painless gene. Genes, Brain, and Behavior, 5(8), 602–613.Find this resource:
Yan, Z., Zhang, W., He, Y., Gorczyca, D., Xiang, Y., Cheng, L.E., Meltzer, S., Jan, L.Y., & Jan, Y.N. (2013). Drosophila NOMPC is a mechanotransduction channel subunit for gentle-touch sensation. Nature, 493, 221–225.Find this resource:
Yemini, E., Jucikas, T., Grundy, L. J., Brown, A. E. X., & Schafer, W. R. (2013). A database of C. elegans behavioral phenotypes. Nature Methods, 10(9), 877–879.Find this resource:
Zahratka, J. A., Williams, P. D. E., Summers, P. J., Komuniecki, R. W., & Bamber, B. A. (2015). Serotonin differentially modulates Ca2+ transients and depolarization in a C. elegans nociceptor. Journal of Neurophysiology, 113(4), 1041.Find this resource:
Zhang, W., Cheng, Li E., Kittelmann, M., Li, J., Petkovic, M., Cheng, T., et al. (2015). Ankyrin repeats convey force to gate the NOMPC mechanotransduction channel. Cell, 162(6), 1391–1403.Find this resource:
Zhang, Y. V., Ni, J., & Montell, C. (2013). The molecular basis for attractive salt taste coding in Drosophila. Science, 340(6138), 1334–1338.Find this resource:
Zhong, L., Bellemer, A., Yan, H., Honjo, K., Robertson, J., Hwang, R. Y., et al. (2012). Thermosensory and non-thermosensory isoforms of Drosophila melanogaster TRPA1 reveal heat sensor domains of a thermoTRP channel. Cell Reports, 1(1), 43–55.Find this resource:
Zhong, L., Hwang, R. Y., & Tracey, W. D. (2010). Pickpocket is a DEG/ENaC protein required for mechanical nociception in Drosophila larvae. Current Biology, 20(5), 429–434.Find this resource: