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date: 17 January 2020

Invertebrate Models of Nociception: Learning from Flies and Worms

Abstract and Keywords

Chronic pain is a significant public health problem, affecting 20–25% of the global population, and there is a clear need for more specific and effective therapeutics. To achieve this, a comprehensive understanding of the underlying mechanisms and molecular machinery driving pain-related diseases is required. The definition of pain as an “unpleasant sensory and emotional experience” associated with tissue injury is innately anthropomorphic, the emotional element being difficult to reconcile in nonhuman organisms. Even simple invertebrates are nevertheless capable of nociception, the neural processing of noxious stimuli. With the significant advantages of simpler nervous systems, experimental tractability, and a high level of conservation, they have a major role to play in advancing our understanding. This chapter reviews our current molecular- and circuit-level understanding of nociception in two of the most widely used invertebrate experimental models, the nematode Caenorhabditis elegans and the fly Drosophila melanogaster. In particular, it summarizes the molecules, cells, and circuits that contribute to nociception in response to diverse noxious stimuli in these model organisms and the behavioral paradigms that we can harness to study them. The chapter discusses how mechanistic insights gained from these experimental systems can improve our understanding of pain in humans.

Keywords: Caenorhabditis elegans, Drosophila melanogaster, nociception, behavior, sensory transduction

Introduction

Chronic pain has an enormous impact on the quality of life of millions of people worldwide, creating a physical and emotional burden to the patients and caregivers affected. Economically, chronic pain is estimated to cost trillions of dollars per year, similar to major human diseases such as cancer, heart disease, or diabetes. It remains difficult to treat, with the majority of patients reporting inadequate pain relief from available therapies. Indeed, the lack of reliable alternative medications is a key contribution to the ongoing opioid epidemic in the United States, which is now spreading globally (Socias & Wood, 2017). Despite a sustained research effort to understand the underlying mechanisms and molecular machinery driving chronic pain, many pain-related diseases remain difficult to treat, in part because we lack a complete understanding of the basic mechanisms driving disease. To this end, developing a better basic understanding of pain biology may inform future treatment in unanticipated ways. Defining the conserved architecture of pain perception across evolution can provide a better understanding of the core mechanisms involved and model organisms thus have an important role in providing basic insight that will lead to new ways to help pain patients. Here we describe our current understanding of the cellular and molecular mechanisms controlling pain/nociception based on genetic studies in the roundworm Caenorhabditis elegans and the fruit fly Drosophila melanogaster.

Pain as defined by the International Association for the Study of Pain is “an unpleasant sensory and emotional experience associated with actual or potential tissue damage, or described in terms of such damage.” The requirement for pain to be “unpleasant” or involve “emotional” experiences is inherently anthropomorphic. By definition this precludes pain from being a conserved process and arguably should be revised based on our current understanding of pain perception across phyla. To avoid this issue, animal researchers often refer to pain perception as nociception, which is defined as “the neural process of encoding and processing noxious stimuli” (Dubin & Patapoutian, 2010, p. 3761). The notion that pain perception is evolutionarily conserved was at one point controversial; however, a multitude of studies have established that pain/nociception is widespread across the animal kingdom (Walters, 2018; Woolf & Walters, 1991). Importantly, molecular studies have demonstrated that the same machinery is used for nociception in animals versus pain in humans. Indeed, both C. elegans and Drosophila studies have been instrumental in identifying key players, such as the Degenerin/Epithelial sodium channel (DEG/ENaC) and transient receptor potential (TRP) channel families. This conservation means that these simple models, with their readily assayable behaviors and genetic amenability, represent huge potential for high-throughput screens, to elucidate mechanisms and identify more effective therapeutics.

Caenorhabditis elegans

With its compact nervous system of only 302 neurons and amenability to genetic, optogenetic, and behavioral studies, C. elegans makes a powerful model in which to better understand neuronal function, at both the cellular and circuit level. Each of the 302 neurons is unique and identifiable, and C. elegans was the first animal to have a completely mapped connectome (B. L. Chen, Hall, & Chklovskii, 2006; White, Southgate, Thomson, & Brenner, 1986). Approximately a third of these neurons are sensory, a third interneurons, and a third motor. Laser ablation has allowed the identification of specific neurons, or groups of neurons, that sense specific noxious stimuli, including mechanical stimulation, temperature, high osmotic strength, heavy metals, and other toxins (Tobin & Bargmann, 2004; Wittenburg & Baumeister, 1999). While some are localized in specialized sensilla at the nose (Fig. 1; two amphids, four cephalic, six inner labial, and six outer labial, all of which sit around the lips), body (two anterior and two posterior deirids), and the tail (two phasmids), others sit below the skin in different regions of the body (Fig. 2).

Invertebrate Models of NociceptionLearning from Flies and Worms

Figure 1. The C. elegans nose contains many sensilla. Schematic illustration showing the locations of the sensilla located around the mouth. The bilaterally symmetrical amphids contain a large number of ciliated sensory neurons, ASE, ASG, ASH, ASI, ASJ, ASK, ADF, ADL, AWA, AWB, AWC, and AFD. CEP, cephalic; IL, inner labial; OL, outer labial.

Invertebrate Models of NociceptionLearning from Flies and Worms

Figure 2. C. elegans nociceptive neurons. (Top) Schematic showing the locations of the sensilla, in which many of the nociceptive neurons are located. The region around the mouth (black box) contains a large number of specialized sensilla, shown in Figure 1. (Middle) The gentle touch neurons are located under the skin and consist of three anterior (a lateral pair, ALM left and right, and AVM, located ventrally) and three posterior (PLM left and right and PVM). (Bottom) The FLP and PVD pairs of neurons are highly branched, also located under the skin, and effectively cover the head region and the remainder of the body, respectively.

Despite its apparent simplicity, C. elegans is nevertheless capable of surprisingly complex sensory perception. It can recognize a wide range of mechanosensory, gustatory, olfactory, and thermal cues and can precisely distinguish concentrations and temperatures, seeking out the specific optimal conditions based on past experience, as well as altering sensory responses as a result of habituation, sensitization, or associative learning using a wide range of environmental and food status indicators. As a result of their unique connectivity, distinct neurons are required for attraction versus repulsion. For example, benzaldehyde is attractive at low concentration and repulsive at high concentration, sensed by distinct pairs of amphid neurons, AWC and ASH, respectively. The neuronal context determines the valence of the sensory cue, and this was particularly elegantly demonstrated by heterologous expression of the diacetyl receptor, ODR-10, which functions in attraction in the AWA pair of amphid neurons. When ODR-10 is expressed in AWC, diacetyl is also attractive, whereas if it is expressed in another amphid neuron, AWB, it becomes repulsive (Troemel, Kimmel, & Bargmann, 1997; Wes & Bargmann, 2001). Appetitive stimuli and some aversive stimuli influence complex behaviors involved in navigation and food seeking, while strongly aversive (nociceptive) stimuli evoke acute escape (nocifensive) responses.

Nociception: Circuits and Molecules Involved in Sensory Transduction

In common with other animals, C. elegans responds to noxious stimuli by initiating a reflex behavior—a reversal followed by a change in direction away from the stimulus. This is a highly reproducible and readily assayed behavior and, while noxious stimuli do elicit other behavioral responses, such as suppression of egg laying (Sanders et al., 2013) and feeding (Bhatla & Horvitz, 2015; Keane & Avery, 2003), as well as longer term changes in behavioral state (Chew et al., 2018), the escape response is by far the most extensively studied. Sinusoidal movement depends on 95 body wall muscle cells along the body, intricately controlled by iterating mini-circuits of motor neurons, with cholinergic A and B neurons providing the excitatory input for a contraction/relaxation wave along the somatic neuromusculature promoting backward or forward movement, respectively. The A and B motorneurons are controlled by two distinct opposing groups of premotor interneurons, with AVA, AVD, and AVE neurons favoring backward movement and AVB and PVC favoring forward movement. The balance of power between these two opposing sets of interneurons thus determines the direction of movement (see D. S. Walker, Chew, & Schafer, 2017, for further explanation). The premotor neurons are the most well-connected neurons in the connectome (Towlson, Vertes, Ahnert, Schafer, & Bullmore, 2013); in particular, they receive direct input from the multitude of nociceptive neurons (Fig. 3). Thus, activation of nociceptors can evoke rapid escape behavior by switching the motor state from forward to backward and triggering a directional reversal. The connectivity here is surprisingly complex, however, with each nociceptor synapsing directly onto a different subset of premotor neurons, using different numbers of chemical and electrical synapses, and outputting indirectly via interneurons which themselves receive input from multiple sensory neurons and output to multiple premotor neurons.

Invertebrate Models of NociceptionLearning from Flies and Worms

Figure 3. C. elegans circuits linking nociceptors to movement. The nociceptors (here represented by three examples, ASH, FLP, and PVD) output directly to the premotor neurons, which constitute two opposing groups, AVA, AVD, and AVE favoring backward movement and AVB and PVC favoring forward movement. These control the cholinergic A and B neurons which provide the excitatory input for a contraction/relaxation wave along the body wall musculature promoting backward or forward movement, respectively. Solid lines indicate chemical synapses; dotted lines indicate gap junctions.

Mechanical Nociception: Distinct Neuron Classes Detect Distinct Types of Stimulus

The first sensory modality to be characterized in detail in C. elegans was mechanosensation, with laser ablation experiments defining the neurons responsible for detecting stimuli of different strengths and at different locations on the body. Strength of stimulus is categorized experimentally as “harsh” or “gentle” touch according to the implement used in behavioral assays—a platinum wire or a hair, respectively (Chalfie, Hart, Rankin, & Goodman, 2014). Both elicit an avoidance response (Chalfie & Sulston, 1981), although the strength of the response (i.e., the distance moved in the escape response) is quantitatively distinct (Li, Kang, Piggott, Feng, & Xu, 2011), and they require distinct groups of neurons. Although the gentle touch neurons are not considered nociceptive, like nociceptors they connect directly to the premotor interneurons and evoke escape behavior when activated.

The gentle touch neurons comprise three anterior neurons, arranged approximately trilaterally (ALM left and right; AVM), and three posterior, arranged similarly (PLM left and right; and a sixth, PVM, whose role in touch sensation is less clear) (Fig. 2). These nonciliated neurons, with unusual 15 protofilament microtubules, extend long neurites along the anterior or posterior body, under the cuticle. Gentle touch to the appropriate region of the body elicits an increase in calcium (Suzuki et al., 2003), while mechanoreceptor currents have been recorded in response to both application and removal of the stimulus (X. Chen & Chalfie, 2015; O’Hagan, Chalfie, & Goodman, 2005). In addition to the directional escape response to gentle touch, the gentle touch neurons also respond to the spatially nonspecific stimulus of substrate vibration, caused by tapping the petri dish, which also causes an escape response (Wicks & Rankin, 1996). The gentle touch neurons make direct connections to the premotor interneurons; both ALM and AVM, for example, have either chemical or electrical (gap junction) synapses to AVD, which is required for the reversal response (Chalfie et al., 1985). They also synapse to additional premotor neurons, though, including those driving forward movement, and it is as yet unclear how the structure of the circuit determines the precise relationship between sensory input and behavioral output.

At least 16 sensory neurons are required for the behavioral response to harsh body touch, acting in distinct body regions (Li et al., 2011; Way & Chalfie, 1989). PVD and FLP are pairs of polymodal multidendritic neurons whose arborizations cover much of the body, and the head and neck, respectively (Fig. 2), and are discussed in more detail later. In anterior harsh touch, FLP acts redundantly with SDQR, AQR, BDU, and ADE while in posterior harsh touch, PVD acts with PDE. For anus touch, PVD and PDE play a minor role, with PHA and PHB being more important. Optogenetic stimulation (by expressing the light-gated Na2+ channel channelrhodopsin under a neuron-specific promoter and using blue light stimulation to activate it) of either PVD or PHA/PHB or FLP or BDU/SDQR/AQR was sufficient to elicit a full behavioral response, underlining their importance (Li et al., 2011). In addition, the so-called gentle touch neurons (ALM, AVM, and PLM) also show calcium responses to harsh touch (distinct from those seen for gentle touch) (Chatzigeorgiou, Yoo, et al., 2010; Suzuki et al., 2003). The functional significance of these in harsh touch is unclear since ablating them alone does not appear to greatly disrupt the behavioral response in the standard platinum wire assay (Li et al., 2011). Comparison of responses to posterior harsh touch following ablation of PVD and PDE provided an insightful observation: While animals still responded to both gentle and harsh touch, the distance moved was the same, suggesting that they are unable to distinguish between the two. This was reflected in the amplitude of calcium transients in the PVC interneurons downstream; the amplitude of responses was the same for the two stimuli, whereas in unablated animals the response to harsh touch is much greater (Li et al., 2011).

Aversive nose touch is also detected by multiple neurons, with ASH and FLP, which we will discuss later, playing a major role. FLP is especially interesting, because it acts within a “mini-circuit” along with the lower threshold mechanosensory neurons OLQ and CEP and the “hub” interneuron RIH. As we have described previously (Chatzigeorgiou & Schafer, 2011; Rabinowitch, Chatzigeorgiou, & Schafer, 2013; Schafer, 2015; D. S. Walker et al., 2017), the electrical synapses via RIH mean that OLQ and CEP can act independently to bring about specific behavioral responses to gentler stimuli (head withdrawal; slowing on food), and also facilitate the harsh touch response of FLP, the only one of the three to directly synapse with the command neurons and thus the vital link for initiation of an escape response.

Thermal Nociception

As with mechanical stimuli, the response to noxious heat depends on the body location of the stimulus; head stimulation results in a reversal and reorientation, tail stimulation produces either a change to, or an acceleration of, forward movement, while mid-body stimulation is probabilistic (Wittenburg & Baumeister, 1999). The amplitude of the response depends on the rate of change, rather than the amplitude of the temperature difference (Mohammadi, Byrne Rodgers, Kotera, & Ryu, 2013). The multidendritic neurons play a central role in noxious heat detection, FLP acting at the head (along with the amphid neuron AFD) and PVD in the mid body, with the phasmid neuron PHC acting in the posterior (Chatzigeorgiou & Schafer, 2011; Liu, Schulze, & Baumeister, 2012; Mohammadi et al., 2013). Interestingly, PVD also responds to, and is required for the behavioral response to, cold shock (Chatzigeorgiou, Yoo, et al., 2010). It is also particularly interesting that AFD, the major sensory neuron responsible for thermotaxis, that is, negotiating temperatures in the range at which worms thrive, also plays a major role in thermo-nociception (Liu et al., 2012). As with the gentle touch neurons and the nose touch circuit, this suggests that a single C. elegans neuron can be both nociceptor and nonnociceptor. This multifunctionality depends on the intricacies of the connectome and the employment of multiple sensor molecules. In the case of AFD, for example, although TAX-2/TAX-4 cyclic nucleotide-gated channels are required for both thermotaxis and thermo-nociception, the guanylyl cyclases are different (GCY-23/GCY-8/GCY-18 in thermotaxis; GCY-12 in nociception); the TRPV receptors OSM-9 and OCR-2 are required only for nociception; and the neurons acting downstream are different (AIY and AIB, respectively) (Liu et al., 2012).

Chemical Nociception

C. elegans responds to a wide range of chemical repellents, including volatiles, high osmotic strength, acid, alkaline, copper, and a range of bitter tastes. ASH plays a major role in avoidance, with other amphid neurons, ADL, ASK, and ASE, playing minor roles that are only evident when ASH is missing (Bargmann, Thomas, & Horvitz, 1990; Hilliard, Bargmann, & Bazzicalupo, 2002; Sambongi et al., 1999, 2000). For quinine avoidance, for example, ASH ablation reduces the response by about 50%, and ASK ablation knocks out a further 25% of the response (the residual response suggesting that there may be other players) (Hilliard, Bergamasco, Arbucci, Plasterk, & Bazzicalupo, 2004). In contrast, ablation of ASH alone completely abolishes the response to high osmotic strength (Hilliard et al., 2004). The response to high protons, on the other hand, is not disrupted by ablation of individual neuron classes; only when pairs of classes ASH, ASK, ADF, and ASE are ablated (Sambongi et al., 2000) is aversive behavior compromised. In both of these examples, it is interesting to note that only ASH is directly connected to the premotor neurons, although all of the neurons synapse extensively with each other, as well as with first-order interneurons. It is worth noting, however, that in assays measuring chemosensory-evoked escape behavior, as opposed to those measuring navigational aversion, the importance of ASH is always prominent (Hilliard et al., 2002).

Polymodal Nociceptors with Distinct Roles

An important challenge in the study of nociception is to understand how polymodal nociceptors can discriminate between multiple types of stimuli, and C. elegans provides us with several experimental models in which to do this.

Located at the muscle–skin interface, the PVD and FLP pairs of sensory neurons envelop the body of the adult worm with highly branched stereotypical arbors of fine sensory processes. Both are polymodal, responding to both mechanical and thermal stimuli, and thus share functional and morphological features with mammalian nociceptors. PVD neurons are born at the second larval stage and extend an anterior-posterior primary dendrite. Secondary dendrites extend perpendicular from this, branching to form T-shaped tertiary dendrites when they reach the outer body wall muscle, from which quaternary dendrites extend to give a menorah-like organization (Fig. 2) (Oren-Suissa, Hall, Treinin, Shemer, & Podbilewicz, 2010; Smith et al., 2010). This precise organization, which has become an important model system in which to dissect the mechanisms of neuronal development, depends on the integration of long- and short-range guidance cues from muscle and skin with cytoskeletal reorganization and branch order-specific transcription factors (see Celestrin, Diaz-Balzac, Tang, Ackley, & Bulow, 2018; O’Brien et al., 2017; Zou et al., 2016, 2018, for more details and references). Additional self-avoidance signals ensure tiling but not overlap of PVD dendrites (Liao, Li, Lee, Chien, & Pan, 2018) and a separate mechanism confines FLP and PVD arbors to separate body regions (Yip & Heiman, 2016). Both FLP and PVD express members of the DEG/ENac and TRP families, which provide clues as to how polymodal functionality is achieved. TRP family members (OSM-9 and OCR-2 in FLP; TRPA-1 in PVD) are implicated in thermosensation (Chatzigeorgiou & Schafer, 2011; Chatzigeorgiou, Yoo, et al., 2010; Liu et al., 2012), while the DEG/ENaCs MEC-10 and DEGT-1 function in harsh touch in PVD (Chatzigeorgiou, Grundy, et al., 2010; Chatzigeorgiou, Yoo, et al., 2010), with FLP also expressing potential candidates, such as DEL-1 (Tavernarakis, Shreffler, Wang, & Driscoll, 1997).

The ASH nociceptors are a pair of ciliated amphid neurons, required for the avoidance response to a wide range of noxious stimuli, including osmotic, heavy metals, SDS, volatile, protons, nose touch, and a wide range of bitter tastes (Bargmann et al., 1990; Hart, Sims, & Kaplan, 1995; Hilliard et al., 2004; Kaplan & Horvitz, 1993; Sambongi et al., 1999, 2000). They have proved a powerful model nociceptor and respond to chemical, osmotic, and mechanical stimulation by initiating a transient increase in global calcium concentration (Hilliard et al., 2005). How can a single neuron class respond to such a diverse range of stimuli, and with such specificity that prolonged exposure to one repellent specifically produces adaptation to this repellent while others are unaffected (Hilliard et al., 2005)? One clue lies in the fact that ASH expresses at least nine different G protein alpha subunits, with distinct roles. We know that one of these, odr-3, is required for responses to all of the chemical repellents tested, for example, whereas another, gpa-3, is required only for quinine, not for copper, SDS, and only slightly for the response to glycerol. Likewise, if we look at downstream signaling components, itr-1, encoding the IP3 receptor, is only required for avoidance of nose touch and benzaldehyde, not for other ASH-mediated avoidance responses (D. S. Walker et al., 2009), providing a suggestion of stimulus-specific pathways.

Like the multidendritic neurons, ASH also coexpresses DEG/ENaCs and TRP channels. As we will see later, the TRPV channels OSM-9 and OCR-2 are required for responses to multiple types of stimuli (Colbert, Smith, & Bargmann, 1997; Hilliard et al., 2005; Tobin et al., 2002), while the role of DEG/ENaC DEG-1 is mechanosensation specific (Geffeney et al., 2011).

In common with ASH, the phasmid neurons PHA and PHB are also polymodal, responding not only to harsh touch (Li et al., 2011) but to a wide range of noxious chemicals (Hilliard et al., 2002; Zou et al., 2017). ASH and PHA/PHA act antagonistically on the motor circuit, integrating sensory information from the head and tail to effectively provide a spatial map of the chemical environment and thus ensure the appropriate escape response. This has been demonstrated for SDS (Hilliard et al., 2002), and it is likely true for other noxious stimuli.

Sensory Transduction Molecules: TRP, DEG/ENaC, and Other Candidates

A particular challenge in elucidating the mechanisms of nociception is identifying the molecules specifically responsible for sensory transduction. In common with Drosophila, C. elegans has played a pivotal role in the discovery of receptor families that we now know to be important across the animal kingdom, and it continues to be a powerful model in which to elucidate their function.

DEG-1, MEC-4, and MEC-10 are founding members of the DEG/ENaC family of amiloride-sensitive sodium channels (DEG for degenerin, as gain-of-function mutations cause degeneration; Chalfie & Au, 1989; Chalfie & Wolinsky, 1990), and they have thus been instrumental in identifying family members in other species. While MEC-4 is essential for gentle touch, MEC-10 plays a lesser role. However, MEC-10 plays an essential role in harsh touch, functioning alongside another DEG/ENaC, DEGT-1, in both the gentle touch neurons and FLP and PVD (Chalfie & Sulston, 1981; Chatzigeorgiou, Grundy, et al., 2010). Likewise in ASH, we know that DEG-1 functions in mechanoreceptor currents, along with at least one other, unidentified, DEG/ENaC (Geffeney et al., 2011). Intriguingly, it also functions in acid avoidance (Y. Wang et al., 2008).

Several TRP family members are also implicated in mechanotransduction. The TRPN channel TRP-4 is required in PDE for posterior harsh touch sensation (L. Kang, Gao, Schafer, Xie, & Xu, 2010), as well as having nonnociceptive mechanosensory roles in CEP (food sensing) and DVA (proprioception) (Kindt, Quast, et al., 2007; Li, Feng, Sternberg, & Xu, 2006). The TRPV channel subunit, OSM-9 (along with OCR-2 or other OCR partners), appears to play a general role, since it is required for multiple sensory modalities (mechanical, thermal, diverse chemicals). In ASH, while it is required for calcium responses to diverse stimuli, including nose touch (Hilliard et al., 2005), it is not required for mechanoreceptor potentials (Geffeney et al., 2011), indicating a role downstream of sensory transduction, perhaps in amplification. TRPA-1 is more of a puzzle; while in OLQ it functions in nose touch, and indeed can confer mechanosensitivity when expressed in CHO cells (Kindt, Viswanath, et al., 2007), in PVD it is dispensable for harsh touch, instead functioning in cold sensation, and confers cold sensitivity when expressed in FLP (Chatzigeorgiou, Yoo, et al., 2010).

As Table 1 shows, the C. elegans genome also encodes members of other protein families that have been implicated in nociception and that have great potential as models for understanding the role of these families in higher animals. Pannexins have been implicated in neuropathic pain (Jeon & Youn, 2015) and UNC-7, a member of the homologous innexin family, functions in gentle and harsh touch, in both the gentle touch neurons and PVD (DSW and WRS, in preparation). Transmembrane channel-like family member TMC-1 functions in ASH in both salt and alkaline avoidance (Chatzigeorgiou, Bang, Hwang, & Schafer, 2013; X. Wang, Li, Liu, Liu, & Xu, 2016). TMC-1 is expressed in several other sensory neurons, and its role there remains to be explored. The role of the C. elegans Piezo ortholog, PEZO-1, remains unknown.

Table 1. C. elegans Ion Channel Family Members Implicated in Diverse Types of Nociception

Amphid

Cephalic

Outer Labial

Deirid

Phasmid

General Body

ASH

AFD

CEP

OLQ

PDE

PHC

TRNs

PVD

FLP

DVA

DEG/ENaCs

MEC-4

Mech

(Chalfie & Sulston, 1981; Chatzigeorgiou, Grundy, et al., 2010)

MEC-10

Mech

(Chalfie & Sulston, 1981; Chatzigeorgiou, Grundy, et al., 2010)

Mech (Chatzigeorgiou, Yoo, et al., 2010)

Mech (Chatzigeorgiou & Schafer, 2011)

DEGT-1

Mech (Chatzigeorgiou, Yoo, et al., 2010)

Mech (Chatzigeorgiou, Yoo, et al., 2010)

DEG-1

Mech (Geffeney et al., 2011)

TRPs

TRP-4 (TRPN)

Mech

(L. Kang et al., 2010; Kindt, Quast, et al., 2007)

Mech

(Li et al., 2011)

Mech

(Li et al., 2006)

TRPA-1 (TRPA)

Mech

(Kindt, Viswanath, et al., 2007)

Cold (Chatzigeorgiou, Yoo, et al., 2010)

OSM-9 (TRPV)

Mech; Chemo (Colbert et al., 1997; Hilliard et al., 2005; Tobin et al., 2002)

Mech (Chatzigeorgiou & Schafer, 2011)

Heat

(Liu et al., 2012)

Heat

(Liu et al., 2012)

OCR-2

Mech; Chemo (Colbert et al., 1997; Tobin et al., 2002)

Heat

(Liu et al., 2012)

Heat

(Liu et al., 2012)

Guanylate Cyclases

GCY-12

Heat

(Liu et al., 2012)

TAX-2

Heat

(Liu et al., 2012)

TAX-4

Heat

(Liu et al., 2012)

Innexins

UNC-7

Mech *

Mech *

TMCs

TMC-1

Salt; Alkali (Chatzigeorgiou et al., 2013; X. Wang et al., 2016)

This table shows the family members, the neurons in which they are known to function, and the nociceptive functions identified. Mech indicates mechanosensation; chem indicates chemosensation.

(*) Note that MEC-4 is implicated only in gentle touch.

(**) DSW and WRS, in preparation.

Many of these examples illustrate the challenges that we face with respect to sensor molecules. First, are they bona fide sensors or merely peripheral players, for which sufficiency, when expressed heterologously, is a good test. But even once we have established this, how can closely related members of a family, or even one specific molecule (such as TRPA-1), perform such different sensory transduction roles?

Modulation of Nociceptor Responses in C. elegans

Nociceptive neurons do not act in isolation but rather receive inputs from multiple other cells. Some of these inputs are in the form of neuromodulators, signaling molecules that effectively change the composition of neuronal circuits by modifying the properties of receiving neurons such that they are recruited to or excluded from an existing circuit (reviewed in Bargmann, 2012; Harris-Warrick & Marder, 1991; Marder, 2012). Neuromodulators, such as neuropeptides or monoamines (in C. elegans: dopamine, serotonin, tyramine, octopamine), primarily signal through G protein–coupled receptors (GPCRs) and can lead to changes in neuronal dynamics and excitability. Neuromodulator functions enable the fixed structure of the wiring diagram to give rise to many distinct circuits by modifying circuit properties or composition, in response to changes in internal and external states.

Here, we will provide some examples of neuromodulation of nociceptor responses to different external stimulations, focusing on the ASH neuron. This is a nonexhaustive list; for more examples, refer to O. Hobert (2003) and Komuniecki, Harris, Hapiak, Wragg, and Bamber (2012).

Neuropeptide Modulation of Nose Touch by Premotor Interneurons

ASH neurons are glutamatergic: They express the vesicular glutamate transporter VGLUT gene eat-4, and eat-4 is required for ASH-mediated escape responses (Berger, Hart, & Kaplan, 1998; Hart, Kass, Shapiro, & Kaplan, 1999). Additionally, the premotor interneurons that are postsynaptic to ASH express the non-NMDA-type glutamate receptor GLR-1, and glr-1 mutants, or mutants with failed GLR-1 clustering at the synapse, are defective in ASH-mediated nose touch behaviors (Hart et al., 1995; Maricq, Peckol, Driscoll, & Bargmann, 1995; Rongo, Whitfield, Rodal, Kim, & Kaplan, 1998). Defects in nose touch behavior in glr-1 mutant animals can be suppressed by mutations in the egl-3 gene encoding a C. elegans ortholog of a proprotein convertase type 2 (PC2). Proprotein convertases like EGL-3 are required for one of the first steps of neuropeptide proprotein processing into bioactive peptides. Mutants of egl-3 are defective in many behaviors, including egg laying, locomotion, and mechanosensation (Jacob & Kaplan, 2003; Kass, Jacob, Kim, & Kaplan, 2001), demonstrating the diverse and important functions of these neuropeptides. How does loss of functional EGL-3 restore ASH-mediated nose touch responses in glr-1 mutants? Behavioral data suggest that this requires glutamate signaling from ASH to premotor interneurons not through GLR-1 (Kass et al., 2001), but through another glutamate receptor, the NMDA-type receptor subunit NMR-1 (Mellem, Brockie, Zheng, Madsen, & Maricq, 2002). Electrophysiological and cell-specific rescue experiments indicate that EGL-3 normally functions in the premotor interneurons to process neuropeptides that act on the ASH neurons, resulting in downregulation of glutamate release from ASH (Kass et al., 2001; Mellem et al., 2002). In an egl-3 mutant, these neuropeptides are absent, so glutamate levels at ASH-premotor interneuron synapses are increased to a level that is sufficient to activate NMR-1 NMDA-type receptors (Mellem et al., 2002). The specific neuropeptides required to modulate the ASH-interneuron circuit are yet to be identified.

In a separate behavioral paradigm, habituation of ASH-mediated escape responses was tested by repeated optogenetic stimulation of the ASH neurons. Mutants lacking GLR-1 rapidly habituate to repeated stimulation of ASH, but like in the case for nose touch, loss of neuropeptide signaling in PC2 (egl-3) mutants and mutants lacking another enzyme required for formation of bioactive peptides (carboxypeptidase E encoded by egl-21) suppressed the glr-1-mediated effect on habituation (Ardiel, Yu, Giles, & Rankin, 2017). A candidate mutant screen revealed that pigment-dispersing factor (PDF) neuropeptides acting through their receptor PDFR-1 in neurons and muscle cells is required for habituation of ASH responses (Ardiel et al., 2017).

Sensory Arousal by Nonlocalized Mechanosensory (Tap) Stimulation

Tapping the plate on which animals are housed results in a nonlocalized mechanical stimulus. After tap, animals first respond with a rapid reversal (escape response, lasting <10 seconds) (Rankin, Beck, & Chiba, 1990), but then change locomotor states to accelerate in forward direction (Chew et al., 2018). This increase in forward locomotion lasts for up to 2 minutes, consistent with the animals being in a state of behavioral arousal. Animals that are in a state of behavioral arousal also display sensory facilitation (Robbins, 1997; van Swinderen & Andretic, 2003). Indeed, if they are first exposed to tap, C. elegans show sensitized ASH-mediated escape responses and enhanced neural activity in the ASH neurons in response to stimulation with an aversive chemical (Chew et al., 2018). This indicates that tap stimulation can result in cross-modal sensory facilitation, where a mechanical stimulus leads to sensitization of chemosensory responses. These locomotor and sensory arousal behaviors require neuropeptides encoded by an FMRFamide-like peptide (flp) gene flp-20, released directly from the mechanosensory gentle touch neurons. These peptides then act on cells expressing their cognate receptor, FRPR-3, where the critical neuron for behavioral arousal is the interneuron RID (Chew et al., 2018). RID is a neuroendocrine cell that does not express classical neurotransmitters or amines (Oliver Hobert, 2013), but it makes many peptides, and its activation potentiates forward movement (Lim et al., 2016). Moreover, optogenetic activation of RID is sufficient to sensitize ASH-mediated escape responses (Chew et al., 2018). These findings support a model in which specific afferent neuropeptides released from mechanosensory neurons convey sensory information to central neuroendocrine interneurons, which in turn may release efferent neuropeptides that convey behavioral state information to peripheral targets, including ASH, to generate behavioral arousal responses that are goal directed and context dependent (Chew et al., 2018). The identity of efferent neuropeptides released from RID (or other neuroendocrine cells) to drive arousal is still unknown, although a role for FLP-14 has been suggested in sustaining the forward motor state (Lim et al., 2016).

Modulation of Chemosensory Responses: The Diverse Roles of Monoamines and Neuropeptides

Volatile Repellents (Odors)

ASH responses to volatile repellents are extensively modulated by monoamines and various neuropeptides (reviewed in Komuniecki et al., 2012). Many studies have focused on the modulation of ASH aversive responses by food or nutritional status. In well-fed conditions, the aversive responses to the repulsive odor 1-octanol are mediated primarily by the ASH neurons (Chao, Komatsu, Fukuto, Dionne, & Hart, 2004) and require glutamatergic signaling (Hart et al., 1999). In contrast, fasting or starvation leads to decreased sensitivity to 1-octanol, and this reduced responsiveness can be rescued by the addition of exogenous serotonin, which therefore appears to mimic the fed state (Chao et al., 2004; G. P. Harris et al., 2009). Serotonin is a monoamine neurotransmitter that is critical for modulation of C. elegans food-associated behaviors (Sawin, Ranganathan, & Horvitz, 2000). The effects of serotonin on ASH aversive responses appear to act via different receptors in different cells: Serotonin acts directly on ASH via a G protein–coupled serotonin receptor SER-5, but also acts on interneurons AIB and AIY via the serotonin-gated ion channel MOD-1 (G. P. Harris et al., 2009; P. D. E. Williams et al., 2018). In addition to serotonin, increasing evidence demonstrates that neuropeptides (G. Harris et al., 2010; Mills et al., 2012) and other monoamines (Ezak & Ferkey, 2010; Wragg et al., 2007) also modulate ASH responses to 1-octanol. For example, octopamine, the invertebrate counterpart to norepinephrine, opposes food/serotonin signals and instead inhibits ASH-mediated aversive responses (Mills et al., 2012; Wragg et al., 2007). Octopamine also acts through multiple receptors in different neurons: It acts directly on ASH neurons via the G protein–coupled octopamine receptor OCTR-1 (Wragg et al., 2007), but also acts on other sensory neurons (ADL, AWB, and ASI) via another GPCR, SER-6 (Mills et al., 2012). Additionally, tyramine, a trace amine with particularly important roles in invertebrates (Roeder, 2005), also opposes serotonin stimulation of ASH aversive responses (Wragg et al., 2007). However, instead of acting directly on ASH, tyramine appears to act primarily on the ASI sensory neurons, also involved in food sensing (Gallagher, Kim, Oldenbroek, Kerr, & You, 2013), via the G protein–coupled tyramine receptor TYRA-3 (Hapiak et al., 2013). Downstream of both octopamine and tyramine, an array of neuropeptides also modulates ASH aversive responses (Komuniecki et al., 2012). For example, the effects of tyramine on ASH-mediated aversive responses require the release of neuropeptides from ASI sensory neurons (Hapiak et al., 2013), whereas octopamine-mediated effects require a distinct set of neuropeptides released from other sensory neurons, including AWB and ADL (Mills et al., 2012). Neuromodulators can therefore modify the composition of the circuit driving ASH-mediated aversive responses to include other sensory or interneurons, in response to nutrition status or other internal/external factors.

Soluble Repellents

Like responses to aversive odors, ASH-mediated aversive behavior in response to soluble chemical repellents such as copper, glycerol, or primaquine can be enhanced by the presence of food. By directly assaying ASH neural activity via calcium imaging, this enhanced behavioral response was found to be a consequence of increased magnitude and duration of ASH sensory responses (Ezcurra, Tanizawa, Swoboda, & Schafer, 2011). In contrast to ASH responses to aversive smells, responses to soluble repellents were not affected by serotonin addition but were robustly increased by exogenous dopamine. Consistent with this, mutants defective in endogenous dopamine signaling (such as cat-2 mutants lacking the tyrosine hydroxylase enzyme required for dopamine biosynthesis) do not show food-dependent enhancement of ASH responses to aversive chemicals. This acute enhancement of ASH responses also requires the DOP-4 dopamine receptor acting cell-autonomously in ASH (Ezcurra et al., 2011).

ASH-mediated escape responses to soluble repellents exhibit adaptation following repeated or prolonged stimulation (Hilliard et al., 2005), which in the presence of food also requires dopamine acting on a different receptor DOP-1, in a cell other than ASH (the interneuron AUA) (Ezcurra et al., 2011; Ezcurra, Walker, Beets, Swoboda, & Schafer, 2016). In a separate behavioral paradigm, habituation of ASH escape responses after repeated optogenetic stimulation also requires dopamine signaling and the DOP-4 receptor acting in a neuron downstream of ASH in the escape circuit (E. L. Ardiel et al., 2016). These findings demonstrate that dopamine modulates ASH responses in an acute or chronic manner by acting through different dopamine receptors in different sensory neurons, including the ASH neurons themselves.

Neuropeptides also modulate ASH adaptation responses to soluble chemicals. The neuropeptide receptors NPR-1 and NPR-2 function in the ASH neurons to promote adaptation in the absence of food. Interestingly, the identified neuropeptide ligands for NPR-1 and NPR-2 (FLP-18 and FLP-21, where FLP-18 only activates NPR-1) are not required for this effect (Ezcurra et al., 2016). This could mean either that another unknown peptide is required, or that a complex interaction between other peptides (and receptors) is taking place. Since neuropeptide signaling increases adaptation of ASH responses in the absence of food, whereas dopamine signaling decreases adaptation in the presence of food, are these modulators acting in the same pathway to modify the adaptation of nociceptor responses? Epistasis experiments indicate that cat-2; npr-1 double mutants act like npr-1 single mutants, suggesting that dopamine signaling acts upstream of NPR-1-mediated neuropeptide signaling to modulate adaptation of ASH responses. Consistent with this, knocking down neuropeptide release from AUA (where the DOP-1 receptor is required for dopamine effects on ASH adaptation behavior, as mentioned earlier) leads to decreased adaptation. These findings suggest a model in which dopaminergic signaling regulates neuropeptide signaling to modulate nociceptive behaviors depending on the fed/nutritional status of the animal (Ezcurra et al., 2016).

Modulatory input from other sensory neurons can also reciprocally affect ASH responses to soluble repellents. Blocking the ASI sensory neurons leads to enhanced ASH responses to copper, suggesting that ASI inhibits ASH nociceptive behavior. This inhibition requires the SER-5 serotonin receptor acting in ASH. ASI neurons are not serotonergic; therefore, the serotonin signal comes from another sensory neuron, ADF. The communication between ASI and ADF to inhibit ASH appears to require not synaptic transmission, but neuropeptide signaling via neuropeptides encoded by nlp-5 and insulin-like peptides ins-1 and ins-3. Interestingly, ASH in turn inhibits ASI responses through octopaminergic signaling acting on the SER-3 octopamine receptor in ASI (M. Guo et al., 2015).

Drosophila melanogaster

Like C. elegans, the fruit fly Drosophila melanogaster has been a potent tool for genetic research, especially with respect to the molecular regulation of the nervous system (Bellen, Tong, & Tsuda, 2010). For example, fly genetics led to the cloning of the first potassium channel shaker (Tempel, Papazian, Schwarz, Jan, & Jan, 1987) and the first identification and cloning of a TRP channel (Montell & Rubin, 1989). Moreover, this system has also been useful for characterizing more complex biological machines, for example the molecular components comprising the circadian clock, a system that is conserved from flies through to humans (Young, 2018). In this section we will discuss specific applications of fruit fly genetics to behaviors in response to noxious stimuli.

Aversive Conditioning

One of the first molecular dissections of fruit fly noxious behavior was through investigations of learning and memory machinery. Odorants and tastants are used to assign nutritional value and determine the potential toxicity of food prior to ingestion (Hallem, Dahanukar, & Carlson, 2006), and long-term memories formed in response to these sensory cues can modify innate responses in naive flies (Tully & Quinn, 1985). Aversive stimuli elicit particularly strong behavioral responses in flies and have long been used for classical learning-dependent conditioning paradigms (Duerr & Quinn, 1982; Mery & Kawecki, 2002; Tully & Quinn, 1985; Wolf et al., 1998). For example, flies exposed to neutral odors concurrently with repeated electric shocks develop a strong aversion to the paired odor, which scales with shock intensity and odor concentration (Tully & Quinn, 1985). These data suggest that flies are highly attuned to noxious stimuli, which they integrate with sensory and nociceptive information to avoid harmful environmental exposures. Considerable effort has been put into defining the molecular factors controlling aversive conditioning in the fly (Keene & Waddell, 2007); however, use of electrical stimulation in this paradigm seems to bypass the sensory nociception machinery.

Subnoxious Thermosensation

In common with other poikilotherms, and as we saw for C. elegans, Drosophila use thermosensation to choose the optimal conditions for growth, and these behaviors are robust and amenable to genetic dissection; however, exposure to extreme temperatures can trigger protective (nocifensive) responses (Barbagallo & Garrity, 2015). The first genetic dissection of thermosensation behavior was performed by the lab of Seymour Benzer at Caltech, where they used a temperature preference assay to identify the anatomical requirements for thermosensation and also describe mutants such as painless that exhibit altered thermosensation behavior (Sayeed & Benzer, 1996; Tracey et al., 2003). Subsequent studies have also implicated members of the TRPA family that we have already encountered in worms, pyrexia (Lee et al., 2005) and TRPA1 (Neely et al., 2011; Zhong et al., 2012), in thermosensation and noxious heat responses. Although certain pyrexia mutants show altered temperature preference, null mutants have normal thermosensation in the innocuous range but exhibit an increased susceptibility to extreme heat defining an acute adaptive response (Lee et al., 2005). Because painless and pyrexia are not evolutionarily conserved in mammals, the highly conserved TRPA1 channel has received intense interest in recent years. TRPA1 was first identified as a warmth-activated ion channel (Viswanath et al., 2003) that is essential for avoidance of hot and cold environments in larvae (Rosenzweig et al., 2005). However, the trpA1 gene encodes thermosensory (TRPA1-A/D) and nonthermosensory (TRPA1-B/C) isoforms with distinct expression patterns that regulate diverse temperature-dependent responses (Zhong et al., 2012). Surprisingly, in the periphery, nonthermosensory TRPA1-C is the only isoform required in larval thermosensory neurons, suggesting that TRPA1 also transmits environmental signals received by other receptors (Zhong et al., 2012). One such pathway is the GQ (Gα49B)-Phospholipase C β (NORPA) signaling axis, which functions upstream of TRPA1 in larval thermotaxis (Kwon, Shim, Wang, & Montell, 2008). Intriguingly, these are the same isoforms of GQ and PLC that are coupled to the light-sensitive rhodopsin GPCR in the eye, and this pathway is indeed also initiated by light-independent functions of rhodopsin, which is expressed at very low levels in thermosensory neurons (Shen et al., 2011). Thus, Drosophila exhibit a robust thermopreference behavior, which is governed by TRPA-family temperature-responsive ion channels.

In adult Drosophila, TRPA1 is required in anterior cell neurons that constitute a warmth-sensor within the adult fly brain to control temperature preference, consistent with a heat-activated biochemical mechanism (Hamada et al., 2008). The central warmth sensor uses the TRPA1-A isoform, which contains an alternatively spliced exon encoding a 37 amino acid intracellular segment that confers temperature sensitivity (Luo, Shen, & Montell, 2017; Zhong et al., 2012). This allows fast temperature changes to trigger behaviors at a lower absolute temperature compared to slower changes (Luo et al., 2017). In contrast to larvae, TRPA1 is not required peripherally in the adult for subnoxious thermosensation, and the TRPA1-C isoform is expressed in gustatory neurons, where it triggers temperature-independent nocifensive responses to electrophilic food (K. Kang et al., 2011). Instead, peripheral heat-sensitive neurons in the fly antenna (Gallio, Ofstad, Macpherson, Wang, & Zuker, 2011) use the gustatory receptor GR28B(D) (Ni et al., 2013). Together, these data suggest that temperatures within the comfortable range are discriminated centrally, whereas noxious heat engages additional pathways for rapid aversive thermotaxis.

In addition to the optimal environment for growth, cold avoidance is critical for the survival of poikilothermic organisms such as Drosophila. In larvae, a distinct set of neurons in the dorsal organ, termed dorsal organ cool cells (DOCCs), are essential for cold avoidance (Klein et al., 2015). These DOCCs use invertebrate-restricted ionotropic receptors (IRs), specifically IR21a and IR25a together with IR93a (Knecht et al., 2016; Ni et al., 2016), to sense and initiate cold aversive behavior. Analogous to the heat-sensing neurons, the adult antenna houses a set of cold cells that sense cold stimuli using TRPP-family receptors brivido1-3 (Gallio et al., 2011). Overall, it is clear that Drosophila use complementary approaches throughout their life cycle to detect and respond to temperature changes in order to maximize growth and survival.

Larval Fly Nociception

The use of Drosophila as a system to investigate the genetics of nociception started with the landmark study by Tracey et al. which showed in response to noxious heat the fruit fly larvae exhibited a robust nociceptive rolling escape response (Tracey, Wilson, Laurent, & Benzer, 2003). Larvae have highly developed systems for detecting noxious temperatures, and thermal nociception can be evaluated using a heated probe (Tracey et al., 2003) or immersion in a warming drop of water (Oswald, Rymarczyk, Chatters, & Sweeney, 2011) to evoke a stereotyped rolling escape response (Fig. 4A). More recently, similar approaches have been applied to study noxious cold using a cooled probe to evoke a full body contraction (CT) (Turner et al., 2016; Turner, Landry, & Galko, 2017) (Fig. 4B). CT can also be quantitatively assessed using a plate-based method to evaluate six to nine larvae at a time, potentially offering higher throughput (Patel & Cox, 2017). In addition to temperature, behavioral responses to mechanical and chemical insults also produce stereotyped escape responses in Drosophila larvae (Im & Galko, 2012; Milinkeviciute, Gentile, & Neely, 2012) (Fig. 4C). Surprisingly, larvae exposed to damaging mechanical or chemical stimuli perform the same rolling escape response that is elicited by noxious heat (Tracey et al., 2003), suggesting that this behavior evolved under selective pressure for one stimulus and was co-opted as a general nocifensive response. In a natural Drosophila population, the majority of larvae can harbor predatory parasitoid wasps (Powell, 1997), and it has been suggested that escaping the initial infection may be an evolutionarily important selective pressure driving nocifensive behavior in the larvae. In fact, in some situations, larvae paradoxically roll toward a noxious stimulus or an attacking wasp, occasionally thwarting the attack and escaping unharmed (Hwang et al., 2007). This mechanical escape response can be experimentally elicited with a calibrated Von Frey filament to deliver a mechanical stimulus (Tracey et al., 2003). Finally, it is emerging that the heat and mechanical nociceptors are also used for avoidance of other hostile conditions. For example, larvae require a lubricating food-derived moisture layer for pupation, which determines their preferred position within a humidity gradient, and optimal positioning within this gradient requires functional nociceptors (Johnson & Carder, 2012). Surprisingly, the same nociceptive cells also detect high-intensity, short-wavelength light, which larvae actively avoid (Xiang et al., 2010). Thus, just as we saw for C. elegans nociceptors, they are polymodal.

Invertebrate Models of NociceptionLearning from Flies and Worms

Figure 4. Nociception assays in Drosophila larvae (A, B, and C) and adults (D and E). (A) Heat avoidance assays. Either a heat probe (46°C) is applied to third instar larvae, which then exhibit a rolling avoidance response, or third instar larvae are placed in a water droplet, which is then heated (29°C) and larvae writhe within the droplet. (B) Cold avoidance assay. Cold probe (10°C) is applied to third instar larvae, which then either contract or lift both anterior and posterior ends or the anterior or posterior end alone. (C) Light touch mechanonociception. Third instar larvae are gently touched on the anterior end; they then contract and turn to avoid the stimulus. (D) Heat escape assay. Adult flies are placed on a plate (25°C), which is then heated (42°C), and flies will jump to avoid the heat. Flies that have defective nociceptive responses fail to jump. (E) Noxious temperature avoidance assay. Adult flies are placed in a chamber at room temperature (25°C). One side of the chamber is heated to noxious temperatures (46°C), and the other to a subnoxious temperature (31°C). Flies with defective nociceptive responses fail to avoid the side with a noxious temperature.

Fly graphic adapted from LES LABORATOIRES SERVIER under Creative Commons Attribution 3.0 unported license: https://creativecommons.org/licenses/by/3.0/.

Pain Circuits in Larvae

A complete understanding of nociception will require integration of genetic and anatomical information to determine how external stimuli are detected and processed to elicit appropriate behavioral responses. The small size and stereotypical wiring patterns of the Drosophila larvae make it particularly well suited to saturated (error-free) network reconstructions (Schneider-Mizell et al., 2016) for systems-level analysis, which can be useful for rapidly discarding incorrect functional hypotheses (Denk, Briggman, & Helmstaedter, 2012). The nociceptive network is extensively remodeled to accommodate the dramatic increase in larval size during development, and it has been proposed that dramatic changes in dendritic structure and connectivity are important for preserving the function of the nociceptive circuits (Gerhard, Andrade, Fetter, Cardona, & Schneider-Mizell, 2017). A key challenge for the field has been deciphering how multimodal nociceptors process different stimuli to generate the appropriate behavior. As discussed earlier, class IV neurons are required for thermal and mechanical nociception (Tracey et al., 2003). Mapping the synaptic outputs of class IV neurons identified direct contacts with A08n and DP-ilp7 neurons (Hu et al., 2017). Interestingly, the DP-ilp7 neurons also receive input from class II and class III neurons (Hu et al., 2017), which detect innocuous touch (Tsubouchi, Caldwell, & Tracey, 2012; Yan et al., 2013). Both A08n and DP-ilp7 neurons are required for the nocifensive rolling response, but only A08n neuronal activation is sufficient for the response, whereas DP-ilp7 neurons play a mechano-specific neuromodulatory role mediated by NPF (the Drosophila NPY homolog) (Hu et al., 2017). In parallel to A08n neurons, activation of the medial cluster of second-order neurons (mCSIs) is also necessary and sufficient for the nocifensive response, at least partly through direct coupling to SNa motor neurons (Yoshino, Morikawa, Hasegawa, & Emoto, 2017). One possibility is that each type of interneuron integrates different costimulatory inputs to tune the output of the nociceptors. For example, DP-ilp7 neurons integrate light touch (Hu et al., 2017), whereas Basin-2/4 interneurons respond to vibration and project onto Goro command neurons that trigger the rolling response (Ohyama et al., 2015). Highlighting the complexity of the intact circuit, A09l (also called Down-and-Back) neurons provide additional connectivity between class 2/3/4 neurons and the Basin interneurons (Burgos et al., 2018). Further integrated ultrastructural and functional mapping of larval pain circuits along with increasingly sophisticated genetic tools for manipulating specific subpopulations will help define the cellular functions of novel nociception genes and decipher why nociceptive processing becomes dysregulated.

In the following sections, we describe the contribution of Drosophila genetics to understanding nociceptive behaviors, first in larvae, followed by insights gained from studies of the adult fly.

Larval Thermal Nociception

Thermal nociception assays are particularly well suited for large-scale screens in Drosophila larvae. The heated probe assay was first used in the genetic screen that identified the temperature-responsive TRPA channel painless, as well as class IV multidendritic sensory neurons, as essential for larval nociception (Tracey et al., 2003). Manipulation of class IV neurons with pickpocket (ppk) GAL4 drivers has shown these cells are necessary for larval heat nociception and optogenetic stimulation is sufficient to elicit the nocifensive rolling response (Hwang et al., 2007). Ppk, which encodes a DEG/ENaC subunit that is important for mechanical nociception, is expressed in all polymodal class IV nociceptors but is not itself required for the response to noxious heat (Zhong, Hwang, & Tracey, 2010), indicating that, as we have already seen in C. elegans PVD neurons, for example, shared polymodal nociceptors use distinct molecular pathways to encode different stimuli. Consistent with its expression in nociceptors and role in thermosensation, TRPA1 is required in larvae for a rapid response to noxious heat (Babcock et al., 2011; Neely et al., 2011; Zhong et al., 2012). Together with Painless and the L-type voltage-gated calcium channel (L-VGCC), TRPA1 triggers Ca2+ influx and a unique nociceptor firing pattern, consisting of high-frequency spike trains and pause periods, in response to a noxious thermal stimulus (Terada et al., 2016). The thermal specificity of the nociceptors is tuned by multiple pathways downstream of Ca2+ influx. The calcium-activated chloride channel Subdued shows a genetic transheterozygous interaction with TRPA1 and Painless (removing one copy of each gene enhances the thermal nociception defect) and likely amplifies TRP channel activity (Jang et al., 2015). Nociceptor responses are also under negative regulation, which is likely important to prevent spontaneous and chronic pain responses without the continued presence of a noxious stimuli. The small conductance Ca2+-activated K+ (SK) channel is responsible for the pauses after spike trains in activated nociceptors (Onodera, Baba, Murakami, Uemura, & Usui, 2017), and knockdown of this channel hypersensitizes larvae to noxious heat (Onodera et al., 2017; Walcott, Mauthner, Tsubouchi, Robertson, & Tracey, 2018). Consistent with the role of Gq and PLCβ signaling in thermosensation discussed earlier, inhibition of this signaling axis reduces basal nociceptive sensitivity (Herman, Willits, & Bellemer, 2018). Different steroid hormones positively and negatively modulate pain sensitivity in vertebrates (Melcangi, Garcia-Segura, & Mensah-Nyagan, 2008). The sole Drosophila steroid hormone ecdysone acting through different isoforms of the ecdysone receptor (EcR) is required for dendritic arborization and sensitivity in class IV neurons (McParland, Follansbee, Vesenka, Panaitiu, & Ganter, 2015). It is unclear how EcR regulates arborization, although mutations in the microtubule-severing protein Katanin p60-like 1 (KAT-60L1) show similar defects, suggesting they may act in the same pathway (Stewart, Tsubouchi, Rolls, Tracey, & Sherwood, 2012). The link between dendritic structure and nociceptor output is further supported by an expression-guided functional analysis of nociceptor enriched genes (Honjo, Mauthner, Wang, Skene, & Tracey, 2016). In many cases, heat insensitivity associated with RNAi knockdown of evolutionarily conserved genes is correlated with severe branching defects, while hyperbranching was only observed in hypersensitive settings (Honjo et al., 2016), highlighting the importance of network structure. While the molecular mechanisms underlying cold nociception have been less intensively investigated, analogous signaling modules have been identified. For example, the TRP channels TRPM, NOMPC, and PKD2 are required in class III neurons for cold-induced CT, and this circuit is functionally dominant when cold- and heat-responsive neurons are coactivated (Turner et al., 2016), providing insight into the hierarchical organization of the nociceptive response.

Larval Mechanical Nociception

Larvae respond to potentially damaging mechanical stimuli using the same heat-responsive class IV neurons to evoke the stereotyped rolling escape response (Hwang et al., 2007). The most commonly used stimulus is a 45 mN von Frey filament, which induces comparable responses in repeated trials of the same individual, indicating that it does not cause permanent damage to the sensory neurons (Kim, Coste, Chadha, Cook, & Patapoutian, 2012). While some of the same molecular machinery is involved in heat and mechanical nociception, such as painless (Tracey et al., 2003), there are also force-specific molecular sensors. Similar to other nociception modalities, there are invertebrate-specific mechanosensing innovations. For example, the first identified Drosophila mechanosensor no mechanoreceptor potential C (nompC) (R. G. Walker, Willingham, & Zuker, 2000), which directly transmits force to the neuronal microtubule network (Zhang et al., 2015), is not conserved in mammals. Similarly, mammalian homologs of the ion channel ppk, whose expression specifically labels class IV sensory neurons (Tracey et al., 2003) and is required for their function (Zhong et al., 2012), have not been functionally confirmed as mechanosensitive channels. In contrast, both fly (Dmpiezo) and mouse (Piezo1 and Piezo2) piezo proteins are sufficient to confer mechanosensitivity to heterologous cells (Coste et al., 2010, 2012). Dmpiezo is specifically required in ppk-positive class IV neurons, where it acts in parallel to ppk1 (ppk) and ppk26 to generate mechanically activated currents (Gorczyca et al., 2014; Y. Guo, Wang, Wang, & Wang, 2014; Kim et al., 2012; Mauthner et al., 2014). Together, knockdown of Dmpiezo and ppk almost entirely eliminate mechanical nociception without having any effect on thermal responses (Kim et al., 2012). Importantly, potentially damaging mechanical stimuli can also be genetically distinguished from light touch sensation, which is used for larval pathfinding. Both painless and Dmpiezo are dispensable for processing gentle touch, which is sensed by the immunoglobulin superfamily protein Turtle (Zhou, Cameron, Chang, & Rao, 2012). Similarly to thermal nociception, modulation of dendritic structure (McParland et al., 2015) and sensory neuron firing patterns (Onodera et al., 2017; Walcott et al., 2018) can induce mechanohypersensitivity.

Adult Fly Nociception

Drosophila nociceptors are extensively remodeled during larval development and persist in the adult fly to sense and respond to noxious stimuli (D. W. Williams & Truman, 2005). Adult flies also exhibit nocifensive escape responses and specifically avoid high temperatures (i.e., above 39°C) (Fig. 4D,E). This preference was first demonstrated using an experimental setup where flies placed on one side of a sealed chamber were attracted to a light source at the far end that was separated by a noxiously heated passage, preventing flies with intact thermal nociception from reaching the light source (Manev & Dimitrijevic, 2004). Additional assays analogous to the standard mammalian hot-plate latency test have been used to confirm the functionality of adult nociceptors and a requirement for painless in the response (Aldrich, Kasuya, Faron, Ishimoto, & Kitamoto, 2010; Xu et al., 2006). The feasibility of novel pain gene discovery in the adult fly was accelerated by the development of a simplified noxious heat avoidance assay where the loss of thermal nociceptive behavior (i.e., analgesia) could be quantified by measuring the percentage of flies that fail to avoid a noxious (46°C) thermal surface when given a safe (31°C) alternative (Neely et al., 2010).

Adult Thermal Nociception

The high-throughput phenotyping of adult nociception enabled unbiased screens using pan-neuronal RNAi knockdown targeting the majority of the Drosophila genome (11,664 genes) (Neely et al., 2010). This approach allowed the discovery of genes that might also have essential developmental functions outside the nervous system and identified 580 candidate thermal nociception genes, the majority of which were conserved in mammals. This functional analysis highlighted the importance of metabolism, proteolysis, and other major signaling pathways in nociceptive behavior. In particular, knockdown of the voltage-gated calcium channel straightjacket (stj) defined a new pain gene with conserved function in mice that was also associated with pain sensitivity in humans (Neely et al., 2010). Secondary bioinformatic analysis of the hits from the genome-wide screen showed that additional genes involved in calcium signaling were overrepresented, including the calcium-permeable TrpA1 channel discussed earlier (Neely et al., 2011). Further systems-level analysis of the 399 candidate genes that have human homologs showed that phosphoinositide signaling constitutes a central node in the pain network and predicted novel mammalian pain genes (PIP5α and PI3Kγ) (Neely et al., 2012). Using experimentally confirmed binding partners, a larger “nociception network” was built that included the allostatin C receptor 1 (AstC-R1), which is related to mammalian pain-modulating opioid receptors (Neely et al., 2012). Functionally, knockdown of either AstC-R1 or its ortholog AstC-R2 hypersensitizes flies to noxious heat (Bachtel, Hovsepian, Nixon, & Eleftherianos, 2018). Further underscoring the plasticity of the nociceptive system, hunger suppresses the behavioral responses to noxious heat in a Leucokinin (Lk)- and Leucokinin receptor (Lkr)-dependent manner (Ohashi & Sakai, 2018). While comparatively few nociception genes have been studied in detail in the adult (versus the larval context), the unbiased nociception networks provide entry points for the investigation of hundreds of conserved components of the molecular machinery underlying the nociceptive response.

Nociceptive Sensitization in Drosophila

Nocifensive responses to noxious stimuli are tightly regulated and poststimulus hypersensitivity is likely an adaptation to protect damaged tissues from further injury during the healing process. If hypersensitivity persists beyond the completion of tissue healing, it can lead to persistent or chronic pain. Nociceptive sensitization can be classified as hyperalgesisa (an exaggerated response to a noxious stimulus) or allodynia (a nocifensive response to a nonnoxious threshold). UV-induced tissue damage was the first sensitization model established in Drosophila larvae. In this system, short-wavelength UVC irradiation is used to damage the larval epidermis prior to evaluating the nociceptive response (Babcock, Landry, & Galko, 2009). In this model, UV irradiation induces allodynia, which requires the apoptotic initiator caspase DRONC in the epidermis, as well as DRONC-independent hyperalgesia that resolves more rapidly. Nociceptive hypersensitivity is likely a direct consequence of tissue damage as it does not appear to require the recruitment of inflammatory cells. TNF signaling is implicated in mammalian pain sensitization (Vogel, Stallforth, & Sommer, 2006), and it is similarly required for the development of UV-induced allodynia. Expression of the TNF ligand Eiger in the UV-damaged epidermis activates its cognate receptor Wengen on class IV neurons, and localized expression of Eiger is sufficient to induce sensitization in the absence of UV treatment (Babcock et al., 2009) (Fig. 5). Surprisingly, despite the involvement of DRONC, UV sensitization does not require apoptotic effector caspases (Jo et al., 2017). Consistent with this genetic data, UV treatment can induce allodynia at doses below the threshold required to trigger apoptosis. Instead, DRONC is required for the localized processing of Eiger and is dispensable if soluble TNF is expressed by class IV neurons, although the interaction is likely indirect since Eiger does not harbor a recognizable caspase cleavage site (Jo et al., 2017). Downstream of TNF/Eiger signaling sensitization is mediated by canonical TNF targets p38/NfκB and the chromatin modulator Enhancer of Zeste (E(z)) (Jo et al., 2017). Hedgehog (Hh) signaling sensitizes nociceptors in parallel to TNF and ectopic activation of both pathways causes more severe allodynia than either pathway alone (Babcock et al., 2011). Interestingly, allodynia is mediated by modulation of distinct TRP channels, with Painless required for Hh- and TNF-mediated allodynia at the lower end of the nociceptive temperature range. In addition, TRPA1-dependent Hh signaling is necessary for UV-induced hyperalgesia (which does not require TNF), and ectopic Hh activity is sufficient to induce hyperalgesia in uninjured tissue (Babcock et al., 2011). Tachykinin signaling through the Tachykinin Receptor (TKR) is required in nociceptors upstream of Hh for UV-induced sensitization (Im et al., 2015). Hh does not likely modulate TRPA1 function directly since BMP/BMPR signaling, which is activated downstream of Hh in other contexts, is also required for allodynia, although the epistatic relationship of these pathways has not yet been explored in nociceptors (Follansbee et al., 2017).

Invertebrate Models of NociceptionLearning from Flies and Worms

Figure 5. Sterile inflammation leads to nociceptor sensitization in Drosophila larvae. Artificially produced UV-C light causes epidermal damage leading to release of the Drosophila TNF ortholog Eiger. Eiger acts on Wengen (TNF receptor ortholog) in type IV sensory neurons, which activates p38/NFκB signaling and causes sensitization. Tachykinin binding to its receptor leads to Hedgehog release through Dispatched, which then acts autocrinally upon the receptor Patched. This leads to further downstream signaling and UV-induced sensitization.

Additional larval models are emerging to explore potential pathophysiological contributions to neuropathic pain, which is a common sequela of chronic diseases such as diabetes mellitus. Diabetes manifests when circulating glucose levels are elevated due to insufficient Insulin ligand or defective signaling through the Insulin Receptor in peripheral tissues. Larvae lacking the Insulin-like Receptor (InR) exhibit normal baseline thermosensation and a typical UV-induced hypersensitivity response as described earlier (Im, Patel, Cox, & Galko, 2018). However, the UV-induced hypersensitivity persists in the InR mutants. As with the TNF/Hh/BMP pathways, InR signaling is required in the nociceptors to modulate thermosensitivity. Interestingly, constitutive activation of InR in class IV neurons prevents hypersensitivity in “diabetic” larvae, suggesting novel potential therapeutic approaches (Im et al., 2018). While larval paradigms have been powerful in informing on the mechanisms regulating nociceptor sensitization, adult fly chronic pain systems are needed to investigate the mechanisms involved in long-term sensitization after injury.

Conserved Genetics of Pain from Invertebrate Model to Human

The high degree of conservation of many cellular functions means that invertebrate models are powerful systems to validate and also predict novel conserved pain genes or pathways (C. Leung, Wilson, Khuong, & Neely, 2013). We have seen how discoveries initially made in worms or flies have informed investigations of mammalian orthologs. The discovery in C. elegans of the role of MEC-4, a founder member of the DEG/ENaC family, in mechanosensation, for example, has ultimately led to extensive investigation of potential mechanosensory roles for its mammalian relatives.

Invertebrate genetics has also provided us with the opportunity to gain important mechanistic details on pain molecules first identified in human studies. For example, pathway modeling of Drosophila pain screening data highlighted a potential role for PIP3 in regulating pain perception, and this led to the appreciation of PI3K gamma as a negative regulator of TRPV1 activation in mammalian DRG (Neely et al., 2012). In some cases it is initially unclear how mutations identified in human pain genes alter pain perception. This is particularly challenging when investigating transcriptional pain nodes. One productive approach used multiple systems, including Drosophila larvae, to decipher disease-associated dysfunction of the PRDM12 transcription factor, which had been associated with hereditary sensory and autonomic neuropathy (HSAN) (Y. C. Chen et al., 2015). Transcriptomic analysis identified a small set of possible target genes whose expression was consistently changed in patient-derived fibroblasts compared to unaffected sex-matched family controls (Nagy et al., 2015). Knockdown of the Drosophila orthologs in class IV neurons revealed novel roles for five possible targets, including tyrotropin-releasing hormone degrading enzyme (TRHDE) in heat nociception (Nagy et al., 2015). Drosophila have also been used to help explain surprising reports of decreased neuropathic pain despite tumor progression in cancer patients administered EGFR inhibitors (Kersten & Cameron, 2012). In mice EGFR inhibitors prevented inflammatory pain with similar potency to morphine, and EGFR heterozygotes were resistant to hypersensitivity from administration of the EGFR ligand epiregulin (EREG) (Martin et al., 2017). Drosophila larvae were used to confirm that the fly ortholog Egfr is specifically required in sensory neurons for thermal nociception (Martin et al., 2017).

Importantly, each conserved gene or pathway potentially represents a new therapeutic target to consider for the management of chronic pain; however, while pain pathways appear highly conserved, it is important to recognize that the mechanisms involved can diverge. For example, screening Drosophila larval nociceptive sensitization implicated the hedgehog pathway, but when tested in mammals, the Hh inhibitor cyclopamine instead prevented tolerance to morphine and enhanced its analgesic activity (Babcock et al., 2011).

Future Perspectives

Thanks to the efforts of hundreds of research groups, and their position at the forefront of technical advances in genetics, optogenetics, and connectomics, the humble fly and worm have become extremely powerful model systems in which to advance our understanding of the cellular and circuit basis of nociception. Of particular significance is the potential for large-scale screens, thanks to the ease with which we can generate large numbers of animals, and the existence of stereotypical behavioral responses to noxious stimuli, which we can use as a readout for nociceptive function. Using automated quantification of these escape responses, we can apply this to high-throughput screening of candidate analgesics, or of mutants that could help to identify further players and thus elucidate the mechanism (see K. Leung, Mohammadi, Ryu, & Nemenman, 2016 and Husson et al., 2012, for examples). This clearly illustrates the potential that invertebrate models hold with respect to future therapeutic developments.

The high degree of conservation of nociceptive mechanisms across the animal kingdom allows us to extrapolate our understanding from these models to the significant challenge of finding more effective human therapeutics for chronic pain. This should help us to resolve some of the significant questions that remain. In particular, while we know the identity of many molecules that play a role in nociception, establishing whether they are actually sensors is more challenging. We have also seen that some of the protein families implicated can have diverse roles in different cellular context, and indeed, individual proteins can have multiple specific functions (C. elegans TRPA-1, for example, acts as a mechanosensor in OLQ, but in PVD it functions not in mechanosensation but in sensing noxious cold). The molecular and cellular basis of this multifunctionality is an important question, and the ability to compare across multiple model animal systems, and to pose questions within the context of a more tractable organism, should play a significant role in resolving these questions.

Author Note:

Equal first author contribution – D. Hesselson and D. S. Walker; equal last author contribution – G. G. Neely and Y. L. Chew.

References

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. doi:10.3109/01677060903419751Find this resource:

Ardiel, E. L., Giles, A. C., Yu, A. J., Lindsay, T. H., Lockery, S. R., & Rankin, C. H. (2016). Dopamine receptor DOP-4 modulates habituation to repetitive photoactivation of a C. elegans polymodal nociceptor. Learning & Memory, 23(10), 495–503. doi:10.1101/lm.041830.116Find this resource:

Ardiel, E. L., Yu, A. J., Giles, A. C., & Rankin, C. H. (2017). Habituation as an adaptive shift in response strategy mediated by neuropeptides. npj Science of Learning, 2(1), 9. doi:10.1038/s41539-017-0011-8Find 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. doi:10.1016/j.cub.2009.03.062Find 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. doi:10.1016/j.cub.2011.08.020Find this resource:

Bachtel, N. D., Hovsepian, G. A., Nixon, D. F., & Eleftherianos, I. (2018). Allatostatin C modulates nociception and immunity in Drosophila. Science Report, 8(1), 7501. doi:10.1038/s41598-018-25855-1Find this resource:

Barbagallo, B., & Garrity, P. A. (2015). Temperature sensation in Drosophila. Current Opinion in Neurobiology, 34, 8–13. doi:10.1016/j.conb.2015.01.002Find this resource:

Bargmann, C. I. (2012). Beyond the connectome: how neuromodulators shape neural circuits. Bioessays, 34(6), 458–465. doi:10.1002/bies.201100185Find 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 Symposium Quantitative Biology, 55, 529–538.Find this resource:

Bellen, H. J., Tong, C., & Tsuda, H. (2010). 100 years of Drosophila research and its impact on vertebrate neuroscience: A history lesson for the future. Nature Reviews Neuroscience, 11(7), 514–522. doi:10.1038/nrn2839Find this resource:

Berger, A. J., Hart, A. C., & Kaplan, J. M. (1998). G alphas-induced neurodegeneration in Caenorhabditis elegans. Journal of Neuroscience, 18(8), 2871–2880.Find this resource:

Bhatla, N., & Horvitz, H. R. (2015). Light and hydrogen peroxide inhibit C. elegans Feeding through gustatory receptor orthologs and pharyngeal neurons. Neuron, 85(4), 804–818. doi:10.1016/j.neuron.2014.12.061Find this resource:

Burgos, A., Honjo, K., Ohyama, T., Qian, C. S., Shin, G. J., Gohl, D. M., … Grueber, W. B. (2018). Nociceptive interneurons control modular motor pathways to promote escape behavior in Drosophila. Elife, 7. doi:10.7554/eLife.26016Find this resource:

Celestrin, K., Diaz-Balzac, C. A., Tang, L. T. H., Ackley, B. D., & Bulow, H. E. (2018). Four specific immunoglobulin domains in UNC-52/Perlecan function with NID-1/Nidogen during dendrite morphogenesis in Caenorhabditis elegans. Development, 145(10). doi:10.1242/dev.158881Find this resource:

Chalfie, M., & Au, M. (1989). Genetic control of differentiation of the Caenorhabditis elegans touch receptor neurons. Science, 243(4894 Pt 1), 1027–1033.Find this resource:

Chalfie, M., Hart, A. C., Rankin, C. H., & Goodman, M. B. (2014). Assaying mechanosensation. In The C. elegans Research Community (Ed.), WormBook. doi:10.1895/wormbook.1.172.1, http://www.wormbook.orgFind this resource:

Chalfie, M., & Sulston, J. (1981). Developmental genetics of the mechanosensory neurons of Caenorhabditis elegans. Developmental Biology, 82(2), 358–370.Find this resource:

Chalfie, M., Sulston, J. E., White, J. G., Southgate, E., Thomson, J. N., & Brenner, S. (1985). The neural circuit for touch sensitivity in Caenorhabditis elegans. Journal of Neuroscience, 5(4), 956–964.Find this resource:

Chalfie, M., & Wolinsky, E. (1990). The identification and suppression of inherited neurodegeneration in Caenorhabditis elegans. Nature, 345(6274), 410–416. doi:10.1038/345410a0Find 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 Science USA, 101(43), 15512–15517. doi:10.1073/pnas.0403369101Find 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(7435), 95–99. doi:10.1038/nature11845Find this resource:

Chatzigeorgiou, M., Grundy, L., Kindt, K. S., Lee, W. H., Driscoll, M., & Schafer, W. R. (2010). Spatial asymmetry in the mechanosensory phenotypes of the C. elegans DEG/ENaC gene mec-10. Journal of Neurophysiology, 104(6), 3334–3344. doi:10.1152/jn.00330.2010Find this resource:

Chatzigeorgiou, M., & Schafer, W. R. (2011). Lateral facilitation between primary mechanosensory neurons controls nose touch perception in C. elegans. Neuron, 70(2), 299–309. doi:10.1016/j.neuron.2011.02.046Find this resource:

Chatzigeorgiou, M., Yoo, S., Watson, J. D., Lee, W. H., Spencer, W. C., Kindt, K. S., … Schafer, W. R. (2010). Specific roles for DEG/ENaC and TRP channels in touch and thermosensation in C. elegans nociceptors. Nature Neuroscience, 13(7), 861–868. doi:10.1038/nn.2581Find this resource:

Chen, B. L., Hall, D. H., & Chklovskii, D. B. (2006). Wiring optimization can relate neuronal structure and function. Proceedings of the National Academy of Science USA, 103(12), 4723–4728. doi:10.1073/pnas.0506806103Find this resource:

Chen, X., & Chalfie, M. (2015). Regulation of mechanosensation in C. elegans through ubiquitination of the MEC-4 mechanotransduction channel. Journal of Neuroscience, 35(5), 2200–2212. doi:10.1523/jneurosci.4082-14.2015Find this resource:

Chen, Y. C., Auer-Grumbach, M., Matsukawa, S., Zitzelsberger, M., Themistocleous, A. C., Strom, T. M., … Senderek, J. (2015). Transcriptional regulator PRDM12 is essential for human pain perception. Nature Genetics, 47(7), 803–808. doi:10.1038/ng.3308Find this resource:

Chew, Y. L., Tanizawa, Y., Cho, Y., Zhao, B., Yu, A. J., Ardiel, E. L., … Schafer, W. R. (2018). An afferent neuropeptide system transmits mechanosensory signals triggering sensitization and arousal in C. elegans. Neuron, 99(6), 1233–1246.e1236. doi:10.1016/j.neuron.2018.08.003Find 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:

Coste, B., Mathur, J., Schmidt, M., Earley, T. J., Ranade, S., Petrus, M. J., … Patapoutian, A. (2010). Piezo1 and Piezo2 are essential components of distinct mechanically activated cation channels. Science, 330(6000), 55–60. doi:10.1126/science.1193270Find this resource:

Coste, B., Xiao, B., Santos, J. S., Syeda, R., Grandl, J., Spencer, K. S., … Patapoutian, A. (2012). Piezo proteins are pore-forming subunits of mechanically activated channels. Nature, 483(7388), 176–181. doi:10.1038/nature10812Find this resource:

Denk, W., Briggman, K. L., & Helmstaedter, M. (2012). Structural neurobiology: Missing link to a mechanistic understanding of neural computation. Nature Reviews Neuroscience, 13(5), 351–358. doi:10.1038/nrn3169Find this resource:

Dubin, A. E., & Patapoutian, A. (2010). Nociceptors: The sensors of the pain pathway. Journal of Clinical Investigation, 120(11), 3760–3772. doi:10.1172/JCI42843Find this resource:

Duerr, J. S., & Quinn, W. G. (1982). Three Drosophila mutations that block associative learning also affect habituation and sensitization. Proceedings of the National Academy of Sciences USA, 79(11), 3646–3650.Find this resource:

Ezak, M. J., & Ferkey, D. M. (2010). The C. elegans D2-like dopamine receptor DOP-3 decreases behavioral sensitivity to the olfactory stimulus 1-octanol. PLoS One, 5(3), e9487. doi:10.1371/journal.pone.0009487Find this resource:

Ezcurra, M., Tanizawa, Y., Swoboda, P., & Schafer, W. R. (2011). Food sensitizes C. elegans avoidance behaviours through acute dopamine signalling. EMBO Journal, 30(6), 1110–1122. doi:10.1038/emboj.2011.22Find 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. doi:10.1523/JNEUROSCI.1128-15.2016Find this resource:

Follansbee, T. L., Gjelsvik, K. J., Brann, C. L., McParland, A. L., Longhurst, C. A., Galko, M. J., & Ganter, G. K. (2017). Drosophila nociceptive sensitization requires BMP signaling via the canonical SMAD pathway. Journal of Neuroscience, 37(35), 8524–8533. doi:10.1523/jneurosci.3458-16.2017Find this resource:

Gallagher, T., Kim, J., Oldenbroek, M., Kerr, R., & You, Y. J. (2013). ASI regulates satiety quiescence in C. elegans. Journal of Neuroscience, 33(23), 9716–9724. doi:10.1523/JNEUROSCI.4493-12.2013Find this resource:

Gallio, M., Ofstad, T. A., Macpherson, L. J., Wang, J. W., & Zuker, C. S. (2011). The coding of temperature in the Drosophila brain. Cell, 144(4), 614–624. doi:10.1016/j.cell.2011.01.028Find this resource:

Geffeney, S. L., Cueva, J. G., Glauser, D. A., Doll, J. C., Lee, T. H., Montoya, M., … Goodman, M. B. (2011). DEG/ENaC but not TRP channels are the major mechanoelectrical transduction channels in a C. elegans nociceptor. Neuron, 71(5), 845–857. doi:10.1016/j.neuron.2011.06.038Find this resource:

Gerhard, S., Andrade, I., Fetter, R. D., Cardona, A., & Schneider-Mizell, C. M. (2017). Conserved neural circuit structure across Drosophila larval development revealed by comparative connectomics. Elife, 6. doi:10.7554/eLife.29089Find this resource:

Gorczyca, D. A., Younger, S., Meltzer, S., Kim, S. E., Cheng, L., Song, W., … Jan, Y. N. (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. doi:10.1016/j.celrep.2014.10.034Find this resource:

Guo, M., Wu, T. H., Song, Y. X., Ge, M. H., Su, C. M., Niu, W. P., … Wu, Z. X. (2015). Reciprocal inhibition between sensory ASH and ASI neurons modulates nociception and avoidance in Caenorhabditis elegans. Nature Communications, 6, 5655. doi:10.1038/ncomms6655Find 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. doi:10.1016/j.celrep.2014.10.020Find this resource:

Hallem, E. A., Dahanukar, A., & Carlson, J. R. (2006). Insect odor and taste receptors. Annual Review of Entomology, 51, 113–135. doi:10.1146/annurev.ento.51.051705.113646Find this resource:

Hamada, F. N., Rosenzweig, M., Kang, K., Pulver, S. R., Ghezzi, A., Jegla, T. J., & Garrity, P. A. (2008). An internal thermal sensor controlling temperature preference in Drosophila. Nature, 454(7201), 217–220. doi:10.1038/nature07001Find this resource:

Hapiak, V., Summers, P., Ortega, A., Law, W. J., Stein, A., & Komuniecki, R. (2013). Neuropeptides amplify and focus the monoaminergic inhibition of nociception in Caenorhabditis elegans. Journal of Neuroscience, 33(35), 14107–14116. doi:10.1523/JNEUROSCI.1324-13.2013Find this resource:

Harris, G., Mills, H., Wragg, R., Hapiak, V., Castelletto, M., Korchnak, A., & Komuniecki, R. W. (2010). The monoaminergic modulation of sensory-mediated aversive responses in Caenorhabditis elegans requires glutamatergic/peptidergic cotransmission. Journal of Neuroscience, 30(23), 7889–7899. doi:10.1523/JNEUROSCI.0497-10.2010Find this resource:

Harris, G. P., Hapiak, V. M., Wragg, R. T., Miller, S. B., Hughes, L. J., Hobson, R. J., … Komuniecki, R. W. (2009). Three distinct amine receptors operating at different levels within the locomotory circuit are each essential for the serotonergic modulation of chemosensation in Caenorhabditis elegans. Journal of Neuroscience, 29(5), 1446–1456. doi:10.1523/JNEUROSCI.4585-08.2009Find this resource:

Harris-Warrick, R. M., & Marder, E. (1991). Modulation of neural networks for behavior. Annual Review of Neuroscience, 14, 39–57. doi:10.1146/annurev.ne.14.030191.000351Find this resource:

Hart, A. C., Kass, J., Shapiro, J. E., & Kaplan, J. M. (1999). Distinct signaling pathways mediate touch and osmosensory responses in a polymodal sensory neuron. Journal of Neuroscience, 19(6), 1952–1958.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. doi:10.1038/378082a0Find this resource:

Herman, J. A., Willits, A. B., & Bellemer, A. (2018). Galphaq and phospholipase Cbeta signaling regulate nociceptor sensitivity in Drosophila melanogaster larvae. Peer Journal, 6, e5632. doi:10.7717/peerj.5632Find 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. doi:10.1038/sj.emboj.7600493Find 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., & 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. doi:10.1038/sj.emboj.7600107Find this resource:

Hobert, O. (2003). Behavioral plasticity in C. elegans: Paradigms, circuits, genes. Journal of Neurobiology, 54(1), 203–223. doi:10.1002/neu.10168Find this resource:

Hobert, O. (2013). The neuronal genome of Caenorhabditis elegans. In The C. elegans Research Community (Ed.), WormBook. doi:10.1895/wormbook.1.161.1, http://www.wormbook.orgFind 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. doi:10.1016/j.celrep.2016.06.003Find this resource:

Hu, C., Petersen, M., Hoyer, N., Spitzweck, B., Tenedini, F., Wang, D., … Soba, P. (2017). Sensory integration and neuromodulatory feedback facilitate Drosophila mechanonociceptive behavior. Nature Neuroscience, 20(8), 1085–1095. doi:10.1038/nn.4580Find this resource:

Husson, S. J., Costa, W. S., Wabnig, S., Stirman, J. N., Watson, J. D., Spencer, W. C., … Gottschalk, A. (2012). Optogenetic analysis of a nociceptor neuron and network reveals ion channels acting downstream of primary sensors. Current Biology, 22(9), 743–752. doi:10.1016/j.cub.2012.02.066Find 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. doi:10.1016/j.cub.2007.11.029Find 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. doi:10.1002/dvdy.22737Find this resource:

Im, S. H., Patel, A. A., Cox, D. N., & Galko, M. J. (2018). Drosophila Insulin receptor regulates the persistence of injury-induced nociceptive sensitization. Disease Models & Mechanisms, 11(5). doi:10.1242/dmm.034231Find 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. doi:10.7554/eLife.10735Find this resource:

Jacob, T. C., & Kaplan, J. M. (2003). The EGL-21 carboxypeptidase E facilitates acetylcholine release at Caenorhabditis elegans neuromuscular junctions. Journal of Neuroscience, 23(6), 2122–2130.Find this resource:

Jang, W., Kim, J. Y., Cui, S., Jo, J., Lee, B. C., Lee, Y., … Kim, C. (2015). The anoctamin family channel subdued mediates thermal nociception in Drosophila. Journal of Biological Chemistry, 290(4), 2521–2528. doi:10.1074/jbc.M114.592758Find this resource:

Jeon, Y. H., & Youn, D. H. (2015). Spinal gap junction channels in neuropathic pain. Korean Journal of Pain, 28(4), 231–235. doi:10.3344/kjp.2015.28.4.231Find this resource:

Jo, J., Im, S. H., Babcock, D. T., Iyer, S. C., Gunawan, F., Cox, D. N., & Galko, M. J. (2017). Drosophila caspase activity is required independently of apoptosis to produce active TNF/Eiger during nociceptive sensitization. Cell Death Disease, 8(5), e2786. doi:10.1038/cddis.2016.474Find this resource:

Johnson, W. A., & Carder, J. W. (2012). Drosophila nociceptors mediate larval aversion to dry surface environments utilizing both the painless TRP channel and the DEG/ENaC subunit, PPK1. PLoS One, 7(3), e32878. doi:10.1371/journal.pone.0032878Find this resource:

Kang, K., Panzano, V. C., Chang, E. C., Ni, L., Dainis, A. M., Jenkins, A. M., … Garrity, P. A. (2011). Modulation of TRPA1 thermal sensitivity enables sensory discrimination in Drosophila. Nature, 481(7379), 76–80. doi:10.1038/nature10715Find this resource:

Kang, L., Gao, J., Schafer, W. R., Xie, Z., & Xu, X. Z. (2010). C. elegans TRP family protein TRP-4 is a pore-forming subunit of a native mechanotransduction channel. Neuron, 67(3), 381–391. doi:10.1016/j.neuron.2010.06.032Find 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:

Kass, J., Jacob, T. C., Kim, P., & Kaplan, J. M. (2001). The EGL-3 proprotein convertase regulates mechanosensory responses of Caenorhabditis elegans. Journal of Neuroscience, 21(23), 9265–9272.Find this resource:

Keane, J., & Avery, L. (2003). Mechanosensory inputs influence Caenorhabditis elegans pharyngeal activity via ivermectin sensitivity genes. Genetics, 164(1), 153–162.Find this resource:

Keene, A. C., & Waddell, S. (2007). Drosophila olfactory memory: single genes to complex neural circuits. Nature Reviews Neuroscience, 8(5), 341–354. doi:10.1038/nrn2098Find this resource:

Kersten, C., & Cameron, M. G. (2012). Cetuximab alleviates neuropathic pain despite tumour progression. BMJ Case Reports, 2012. doi:10.1136/bcr.12.2011.5374Find 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. doi:10.1038/nature10801Find this resource:

Kindt, K. S., Quast, K. B., Giles, A. C., De, S., Hendrey, D., Nicastro, I., … Schafer, W. R. (2007). Dopamine mediates context-dependent modulation of sensory plasticity in C. elegans. Neuron, 55(4), 662–676. doi:10.1016/j.neuron.2007.07.023Find this resource:

Kindt, K. S., Viswanath, V., Macpherson, L., Quast, K., Hu, H., Patapoutian, A., & Schafer, W. R. (2007). Caenorhabditis elegans TRPA-1 functions in mechanosensation. Nature Neuroscience, 10(5), 568–577. doi:10.1038/nn1886Find this resource:

Klein, M., Afonso, B., Vonner, A. J., Hernandez-Nunez, L., Berck, M., Tabone, C. J., … Samuel, A. D. (2015). Sensory determinants of behavioral dynamics in Drosophila thermotaxis. Proceedings of the National Academy of Sciences USA, 112(2), E220–E229. doi:10.1073/pnas.1416212112Find this resource:

Knecht, Z. A., Silbering, A. F., Ni, L., Klein, M., Budelli, G., Bell, R., … Garrity, P. A. (2016). Distinct combinations of variant ionotropic glutamate receptors mediate thermosensation and hygrosensation in Drosophila. Elife, 5. doi:10.7554/eLife.17879Find this resource:

Komuniecki, R., Harris, G., Hapiak, V., Wragg, R., & Bamber, B. (2012). Monoamines activate neuropeptide signaling cascades to modulate nociception in C. elegans: A useful model for the modulation of chronic pain? Invertebrate Neuroscience, 12(1), 53–61. doi:10.1007/s10158-011-0127-0Find this resource:

Kwon, Y., Shim, H. S., Wang, X., & Montell, C. (2008). Control of thermotactic behavior via coupling of a TRP channel to a phospholipase C signaling cascade. Nature Neuroscience, 11(8), 871–873. doi:10.1038/nn.2170Find this resource:

Lee, Y., Lee, Y., Lee, J., Bang, S., Hyun, S., Kang, J., … Kim, J. (2005). Pyrexia is a new thermal transient receptor potential channel endowing tolerance to high temperatures in Drosophila melanogaster. Nature Genetics, 37(3), 305–310. doi:10.1038/ng1513Find this resource:

Leung, C., Wilson, Y., Khuong, T. M., & Neely, G. G. (2013). Fruit flies as a powerful model to drive or validate pain genomics efforts. Pharmacogenomics, 14(15), 1879–1887. doi:10.2217/pgs.13.196Find this resource:

Leung, K., Mohammadi, A., Ryu, W. S., & Nemenman, I. (2016). Stereotypical escape behavior in Caenorhabditis elegans allows quantification of effective heat stimulus level. PLoS Computational Biology, 12(12), e1005262. doi:10.1371/journal.pcbi.1005262Find this resource:

Li, W., Feng, Z., Sternberg, P. W., & Xu, X. Z. (2006). A C. elegans stretch receptor neuron revealed by a mechanosensitive TRP channel homologue. Nature, 440(7084), 684–687. doi:10.1038/nature04538Find 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 Caenorhabditis elegans. Nature Communications, 2, 315. doi:10.1038/ncomms1308Find this resource:

Liao, C. P., Li, H., Lee, H. H., Chien, C. T., & Pan, C. L. (2018). Cell-autonomous regulation of dendrite self-avoidance by the Wnt secretory factor MIG-14/Wntless. Neuron, 98(2), 320–334.e326. doi:10.1016/j.neuron.2018.03.031Find this resource:

Lim, M. A., Chitturi, J., Laskova, V., Meng, J., Findeis, D., Wiekenberg, A., … Zhen, M. (2016). Neuroendocrine modulation sustains the C. elegans forward motor state. Elife, 5. doi:10.7554/eLife.19887Find 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. doi:10.1371/journal.pone.0032360Find 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(1), 34–41. doi:10.1038/nn.4416Find this resource:

Manev, H., & Dimitrijevic, N. (2004). Drosophila model for in vivo pharmacological analgesia research. European Journal of Pharmacology, 491(2–3), 207–208. doi:10.1016/j.ejphar.2004.03.030Find this resource:

Marder, E. (2012). Neuromodulation of neuronal circuits: Back to the future. Neuron, 76(1), 1–11. doi:10.1016/j.neuron.2012.09.010Find 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. doi:10.1038/378078a0Find this resource:

Martin, L. J., Smith, S. B., Khoutorsky, A., Magnussen, C. A., Samoshkin, A., Sorge, R. E., … Diatchenko, L. (2017). Epiregulin and EGFR interactions are involved in pain processing. Journal of Clinical Investigation, 127(9), 3353–3366. doi:10.1172/jci87406Find this resource:

Mauthner, S. E., Hwang, R. Y., Lewis, A. H., Xiao, Q., Tsubouchi, A., Wang, Y., … Tracey, W. D., Jr. (2014). Balboa binds to pickpocket in vivo and is required for mechanical nociception in Drosophila larvae. Current Biology, 24(24), 2920–2925. doi:10.1016/j.cub.2014.10.038Find 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. doi:10.1371/journal.pone.0140785Find this resource:

Melcangi, R. C., Garcia-Segura, L. M., & Mensah-Nyagan, A. G. (2008). Neuroactive steroids: State of the art and new perspectives. Cell and Molecular Life Sciences, 65(5), 777–797. doi:10.1007/s00018-007-7403-5Find 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:

Mery, F., & Kawecki, T. J. (2002). Experimental evolution of learning ability in fruit flies. Proceedings of the National Academy of Sciences USA, 99(22), 14274–14279. doi:10.1073/pnas.222371199Find this resource:

Milinkeviciute, G., Gentile, C., & Neely, G. G. (2012). Drosophila as a tool for studying the conserved genetics of pain. Clinical Genetics, 82(4), 359–366. doi:10.1111/j.1399-0004.2012.01941.xFind this resource:

Mills, H., Wragg, R., Hapiak, V., Castelletto, M., Zahratka, J., Harris, G., … Komuniecki, R. (2012). Monoamines and neuropeptides interact to inhibit aversive behaviour in Caenorhabditis elegans. EMBO Journal, 31(3), 667–678. doi:10.1038/emboj.2011.422Find 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, 66. doi:10.1186/1471-2202-14-66Find this resource:

Montell, C., & Rubin, G. M. (1989). Molecular characterization of the Drosophila trp locus: A putative integral membrane protein required for phototransduction. Neuron, 2(4), 1313–1323.Find this resource:

Nagy, V., Cole, T., Van Campenhout, C., Khoung, T. M., Leung, C., Vermeiren, S., … Penninger, J. M. (2015). The evolutionarily conserved transcription factor PRDM12 controls sensory neuron development and pain perception. Cell Cycle, 14(12), 1799–1808. doi:10.1080/15384101.2015.1036209Find this resource:

Neely, G. G., Hess, A., Costigan, M., Keene, A. C., Goulas, S., Langeslag, M., … Penninger, J. M. (2010). A genome-wide Drosophila screen for heat nociception identifies alpha2delta3 as an evolutionarily conserved pain gene. Cell, 143(4), 628–638. doi:10.1016/j.cell.2010.09.047Find this resource:

Neely, G. G., Keene, A. C., Duchek, P., Chang, E. C., Wang, Q. P., Aksoy, Y. A., … Penninger, J. M. (2011). TrpA1 regulates thermal nociception in Drosophila. PLoS One, 6(8), e24343. doi:10.1371/journal.pone.0024343Find this resource:

Neely, G. G., Rao, S., Costigan, M., Mair, N., Racz, I., Milinkeviciute, G., … Penninger, J. M. (2012). Construction of a global pain systems network highlights phospholipid signaling as a regulator of heat nociception. PLoS Genetics, 8(12), e1003071. doi:10.1371/journal.pgen.1003071Find this resource:

Ni, L., Bronk, P., Chang, E. C., Lowell, A. M., Flam, J. O., Panzano, V. C., … Garrity, P. A. (2013). A gustatory receptor paralogue controls rapid warmth avoidance in Drosophila. Nature, 500(7464), 580–584. doi:10.1038/nature12390Find this resource:

Ni, L., Klein, M., Svec, K. V., Budelli, G., Chang, E. C., Ferrer, A. J., … Garrity, P. A. (2016). The ionotropic receptors IR21a and IR25a mediate cool sensing in Drosophila. Elife, 5. doi:10.7554/eLife.13254Find this resource:

O’Brien, B. M. J., Palumbos, S. D., Novakovic, M., Shang, X., Sundararajan, L., & Miller, D. M., 3rd. (2017). Separate transcriptionally regulated pathways specify distinct classes of sister dendrites in a nociceptive neuron. Developmental Biology, 432(2), 248–257. doi:10.1016/j.ydbio.2017.10.009Find 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. doi:10.1038/nn1362Find this resource:

Ohashi, H., & Sakai, T. (2018). Leucokinin signaling regulates hunger-driven reduction of behavioral responses to noxious heat in Drosophila. Biochemical and Biophysical Research Communications, 499(2), 221–226. doi:10.1016/j.bbrc.2018.03.132Find this resource:

Ohyama, T., Schneider-Mizell, C. M., Fetter, R. D., Aleman, J. V., Franconville, R., Rivera-Alba, M., … Zlatic, M. (2015). A multilevel multimodal circuit enhances action selection in Drosophila. Nature, 520(7549), 633–639. doi:10.1038/nature14297Find this resource:

Onodera, K., Baba, S., Murakami, A., Uemura, T., & Usui, T. (2017). Small conductance Ca(2+)-activated K(+) channels induce the firing pause periods during the activation of Drosophila nociceptive neurons. Elife, 6. doi:10.7554/eLife.29754Find this resource:

Oren-Suissa, M., Hall, D. H., Treinin, M., Shemer, G., & Podbilewicz, B. (2010). The fusogen EFF-1 controls sculpting of mechanosensory dendrites. Science, 328(5983), 1285–1288. doi:10.1126/science.1189095Find this resource:

Oswald, M., Rymarczyk, B., Chatters, A., & Sweeney, S. T. (2011). A novel thermosensitive escape behavior in Drosophila larvae. Fly (Austin), 5(4), 304–306. doi:10.4161/fly.5.4.17810Find this resource:

Patel, A. A., & Cox, D. N. (2017). Behavioral and functional assays for investigating mechanisms of noxious cold detection and multimodal sensory processing in Drosophila larvae. Bio Protocol, 7(13). doi:10.21769/BioProtoc.2388Find this resource:

Powell, J. R. (1997). Progress and prospects in evolutionary biology: The Drosophila model. New York, NY: Oxford University Press.Find this resource:

Rabinowitch, I., Chatzigeorgiou, M., & Schafer, W. R. (2013). A gap junction circuit enhances processing of coincident mechanosensory inputs. Current Biology, 23(11), 963–967. doi:10.1016/j.cub.2013.04.030Find this resource:

Rankin, C. H., Beck, C. D., & Chiba, C. M. (1990). Caenorhabditis elegans: A new model system for the study of learning and memory. Behavior Brain Research, 37(1), 89–92.Find this resource:

Robbins, T. W. (1997). Arousal systems and attentional processes. Biological Psychology, 45(1–3), 57–71.Find this resource:

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

Rongo, C., Whitfield, C. W., Rodal, A., Kim, S. K., & Kaplan, J. M. (1998). LIN-10 is a shared component of the polarized protein localization pathways in neurons and epithelia. Cell, 94(6), 751–759.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. doi:10.1101/gad.1278205Find 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:

Sambongi, Y., Takeda, K., Wakabayashi, T., Ueda, I., Wada, Y., & Futai, M. (2000). Caenorhabditis elegans senses protons through amphid chemosensory neurons: Proton signals elicit avoidance behavior. Neuroreport, 11(10), 2229–2232.Find this resource:

Sanders, J., Nagy, S., Fetterman, G., Wright, C., Treinin, M., & Biron, D. (2013). The Caenorhabditis elegans interneuron ALA is (also) a high-threshold mechanosensor. BMC Neuroscience, 14, 156. doi:10.1186/1471-2202-14-156Find this resource:

Sawin, E. R., Ranganathan, R., & Horvitz, H. R. (2000). C. elegans locomotory rate is modulated by the environment through a dopaminergic pathway and by experience through a serotonergic pathway. Neuron, 26(3), 619–631.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:

Schafer, W. R. (2015). Mechanosensory molecules and circuits in C. elegans. Pflugers Archives, 467(1), 39–48. doi:10.1007/s00424-014-1574-3Find this resource:

Schneider-Mizell, C. M., Gerhard, S., Longair, M., Kazimiers, T., Li, F., Zwart, M. F., … Cardona, A. (2016). Quantitative neuroanatomy for connectomics in Drosophila. Elife, 5. doi:10.7554/eLife.12059Find this resource:

Shen, W. L., Kwon, Y., Adegbola, A. A., Luo, J., Chess, A., & Montell, C. (2011). Function of rhodopsin in temperature discrimination in Drosophila. Science, 331(6022), 1333–1336. doi:10.1126/science.1198904Find this resource:

Smith, C. J., Watson, J. D., Spencer, W. C., O’Brien, T., Cha, B., Albeg, A., … Miller, D. M., 3rd. (2010). Time-lapse imaging and cell-specific expression profiling reveal dynamic branching and molecular determinants of a multi-dendritic nociceptor in C. elegans. Developmental Biology, 345(1), 18–33. doi:10.1016/j.ydbio.2010.05.502Find this resource:

Socias, M. E., & Wood, E. (2017). Epidemic of deaths from fentanyl overdose. British Medical Journal, 358, j4355. doi:10.1136/bmj.j4355Find this resource:

Stewart, A., Tsubouchi, A., Rolls, M. M., Tracey, W. D., & Sherwood, N. T. (2012). Katanin p60-like1 promotes microtubule growth and terminal dendrite stability in the larval class IV sensory neurons of Drosophila. Journal of Neuroscience, 32(34), 11631–11642. doi:10.1523/jneurosci.0729-12.2012Find this resource:

Suzuki, H., Kerr, R., Bianchi, L., Frokjaer-Jensen, C., Slone, D., Xue, J., … Schafer, W. R. (2003). In vivo imaging of C. elegans mechanosensory neurons demonstrates a specific role for the MEC-4 channel in the process of gentle touch sensation. Neuron, 39(6), 1005–1017.Find this resource:

Tavernarakis, N., Shreffler, W., Wang, S., & Driscoll, M. (1997). unc-8, a DEG/ENaC family member, encodes a subunit of a candidate mechanically gated channel that modulates C. elegans locomotion. Neuron, 18(1), 107–119.Find this resource:

Tempel, B. L., Papazian, D. M., Schwarz, T. L., Jan, Y. N., & Jan, L. Y. (1987). Sequence of a probable potassium channel component encoded at Shaker locus of Drosophila. Science, 237(4816), 770–775.Find this resource:

Terada, S., 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. doi:10.7554/eLife.12959Find this resource:

Tobin, D. M., & Bargmann, C. I. (2004). Invertebrate nociception: Behaviors, neurons and molecules. Journal of Neurobiology, 61(1), 161–174. doi:10.1002/neu.20082Find this resource:

Tobin, D. M., Madsen, D. M., Kahn-Kirby, A., Peckol, E. L., Moulder, G., Barstead, R., … Bargmann, C. I. (2002). Combinatorial expression of TRPV channel proteins defines their sensory functions and subcellular localization in C. elegans neurons. Neuron, 35(2), 307–318.Find this resource:

Towlson, E. K., Vertes, P. E., Ahnert, S. E., Schafer, W. R., & Bullmore, E. T. (2013). The rich club of the C. elegans neuronal connectome. Journal of Neuroscience, 33(15), 6380–6387. doi:10.1523/JNEUROSCI.3784-12.2013Find 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:

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:

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(22), 2124–2134. doi:10.1016/j.cub.2012.09.019Find this resource:

Tully, T., & Quinn, W. G. (1985). Classical conditioning and retention in normal and mutant Drosophila melanogaster. Journal of Comparative Physiology A, 157(2), 263–277.Find this resource:

Turner, H. N., Armengol, K., Patel, A. A., Himmel, N. J., Sullivan, L., Iyer, S. C., … Cox, D. N. (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(23), 3116–3128. doi:10.1016/j.cub.2016.09.038Find this resource:

Turner, H. N., Landry, C., & Galko, M. J. (2017). Novel assay for cold nociception in Drosophila larvae. Journal of Visualized Experiments, 122. doi:10.3791/55568Find this resource:

van Swinderen, B., & Andretic, R. (2003). Arousal in Drosophila. Behavioural Processes, 64(2), 133–144. doi:10.1016/s0376-6357(03)00131-1Find this resource:

Viswanath, V., Story, G. M., Peier, A. M., Petrus, M. J., Lee, V. M., Hwang, S. W., … Jegla, T. (2003). Opposite thermosensor in fruitfly and mouse. Nature, 423(6942), 822–823. doi:10.1038/423822aFind this resource:

Vogel, C., Stallforth, S., & Sommer, C. (2006). Altered pain behavior and regeneration after nerve injury in TNF receptor deficient mice. Journal of the Peripheral Nervous System, 11(4), 294–303. doi:10.1111/j.1529-8027.2006.00101.xFind this resource:

Walcott, K. C. E., Mauthner, S. E., Tsubouchi, A., Robertson, J., & Tracey, W. D. (2018). The Drosophila small conductance calcium-activated potassium channel negatively regulates nociception. Cell Reports, 24(12), 3125–3132.e3123. doi:10.1016/j.celrep.2018.08.070Find this resource:

Walker, D. S., Chew, Y. L., & Schafer, W. R. (2017). Genetics of behavior in C. elegans. In John H. Byrne (Ed.), The Oxford handbook of invertebrate neurobiology. New York: Oxford University Press, 151–170. doi:10.1093/oxfordhb/9780190456757.013.5Find this resource:

Walker, D. S., Vazquez-Manrique, R. P., Gower, N. J., 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), e1000636. doi:10.1371/journal.pgen.1000636Find this resource:

Walker, R. G., Willingham, A. T., & Zuker, C. S. (2000). A Drosophila mechanosensory transduction channel. Science, 287(5461), 2229–2234.Find this resource:

Walters, E. T. (2018). Nociceptive biology of molluscs and arthropods: Evolutionary clues about functions and mechanisms potentially related to pain. Frontiers in Physiology, 9, 1049. doi:10.3389/fphys.2018.01049Find this resource:

Wang, X., Li, G., Liu, J., Liu, J., & Xu, X. Z. (2016). TMC-1 mediates alkaline sensation in C. elegans through nociceptive neurons. Neuron, 91(1), 146–154. doi:10.1016/j.neuron.2016.05.023Find this resource:

Wang, Y., Apicella, A., Jr., Lee, S. K., Ezcurra, M., Slone, R. D., Goldmit, M., … Bianchi, L. (2008). A glial DEG/ENaC channel functions with neuronal channel DEG-1 to mediate specific sensory functions in C. elegans. EMBO Journal, 27(18), 2388–2399. doi:10.1038/emboj.2008.161Find 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:

Wes, P. D., & Bargmann, C. I. (2001). C. elegans odour discrimination requires asymmetric diversity in olfactory neurons. Nature, 410(6829), 698–701. doi:10.1038/35070581Find 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., & Rankin, C. H. (1996). The integration of antagonistic reflexes revealed by laser ablation of identified neurons determines habituation kinetics of the Caenorhabditis elegans tap withdrawal response. Journal of Comparative Physiology A, 179(5), 675–685.Find this resource:

Williams, D. W., & Truman, J. W. (2005). Cellular mechanisms of dendrite pruning in Drosophila: insights from in vivo time-lapse of remodeling dendritic arborizing sensory neurons. Development, 132(16), 3631–3642. doi:10.1242/dev.01928Find this resource:

Williams, P. D. E., Zahratka, J. A., Rodenbeck, M., Wanamaker, J., Linzie, H., & Bamber, B. A. (2018). Serotonin disinhibits a Caenorhabditis elegans sensory neuron by suppressing Ca(2+)-dependent negative feedback. Journal of Neuroscience, 38(8), 2069–2080. doi:10.1523/JNEUROSCI.1908-17.2018Find 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:

Wolf, R., Wittig, T., Liu, L., Wustmann, G., Eyding, D., & Heisenberg, M. (1998). Drosophila mushroom bodies are dispensable for visual, tactile, and motor learning. Learning & Memory, 5(1–2), 166–178.Find this resource:

Woolf, C. J., & Walters, E. T. (1991). Common patterns of plasticity contributing to nociceptive sensitization in mammals and Aplysia. Trends in Neuroscience, 14(2), 74–78.Find this resource:

Wragg, R. T., Hapiak, V., Miller, S. B., Harris, G. P., Gray, J., Komuniecki, P. R., & Komuniecki, R. W. (2007). Tyramine and octopamine independently inhibit serotonin-stimulated aversive behaviors in Caenorhabditis elegans through two novel amine receptors. Journal of Neuroscience, 27(49), 13402–13412. doi:10.1523/JNEUROSCI.3495-07.2007Find 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. doi:10.1038/nature09576Find 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. doi:10.1111/j.1601-183X.2006.00213.xFind this resource:

Yan, Z., Zhang, W., He, Y., Gorczyca, D., Xiang, Y., Cheng, L. E., … Jan, Y. N. (2013). Drosophila NOMPC is a mechanotransduction channel subunit for gentle-touch sensation. Nature, 493(7431), 221–225. doi:10.1038/nature11685Find this resource:

Yip, Z. C., & Heiman, M. G. (2016). Duplication of a single neuron in C. elegans reveals a pathway for dendrite tiling by mutual repulsion. Cell Reports, 15(10), 2109–2117. doi:10.1016/j.celrep.2016.05.003Find this resource:

Yoshino, J., Morikawa, R. K., Hasegawa, E., & Emoto, K. (2017). Neural circuitry that evokes escape behavior upon activation of nociceptive sensory neurons in Drosophila larvae. Current Biology, 27(16), 2499–2504.e2493. doi:10.1016/j.cub.2017.06.068Find this resource:

Young, M. W. (2018). Time travels: A 40-year journey from Drosophila’s Clock mutants to human circadian disorders (Nobel lecture). Angewandte Chemie International Edition, 57(36), 11532–11539. doi:10.1002/anie.201803337Find this resource:

Zhang, W., Cheng, L. E., Kittelmann, M., Li, J., Petkovic, M., Cheng, T., … Jan, Y. N. (2015). Ankyrin repeats convey force to gate the NOMPC mechanotransduction channel. Cell, 162(6), 1391–1403. doi:10.1016/j.cell.2015.08.024Find this resource:

Zhong, L., Bellemer, A., Yan, H., Ken, H., Jessica, R., Hwang, R. Y., … Tracey, W. D. (2012). Thermosensory and nonthermosensory isoforms of Drosophila melanogaster TRPA1 reveal heat-sensor domains of a thermoTRP Channel. Cell Reports, 1(1), 43–55. doi:10.1016/j.celrep.2011.11.002Find 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. doi:10.1016/j.cub.2009.12.057Find this resource:

Zhou, Y., Cameron, S., Chang, W. T., & Rao, Y. (2012). Control of directional change after mechanical stimulation in Drosophila. Molecular Brain, 5, 39. doi:10.1186/1756-6606-5-39Find this resource:

Zou, W., Cheng, H., Li, S., Yue, X., Xue, Y., Chen, S., & Kang, L. (2017). Polymodal responses in C. elegans phasmid neurons rely on multiple intracellular and intercellular signaling pathways. Scientific Reports, 7, 42295. doi:10.1038/srep42295Find this resource:

Zou, W., Dong, X., Broederdorf, T. R., Shen, A., Kramer, D. A., Shi, R., … Shen, K. (2018). A dendritic guidance receptor complex brings together distinct actin regulators to drive efficient F-actin assembly and branching. Developmental Cell, 45(3), 362–375.e363. doi:10.1016/j.devcel.2018.04.008Find this resource:

Zou, W., Shen, A., Dong, X., Tugizova, M., Xiang, Y. K., & Shen, K. (2016). A multi-protein receptor-ligand complex underlies combinatorial dendrite guidance choices in C. elegans. Elife, 5. doi:10.7554/eLife.18345Find this resource: