Introduction

The ability to detect tissue-damaging noxious stimuli and mount an escape response is essential for survival. Likewise, prolonged hypersensitivity following injury is an important form of plasticity that protects an animal from further damage. In Drosophila, a single class of identified somatosensory neurons (SSNs), class IV dendrite arborization (C4da) neurons, are necessary and sufficient for nociception; inactivating C4da neurons renders larvae insensitive to noxious stimuli whereas activating these neurons drives nocifensive behavior responses (Hwang et al., 2007; Hu et al., 2017; Burgos et al., 2018). A variety of agents that cause tissue damage including UV irradiation and chemical toxins induce long-lasting allodynia and hyperalgesia (Babcock et al., 2009; Boiko et al., 2017), but this damage-induced hypersensitivity develops on a timescale of hours. Drosophila also display acute hypersensitivity noxious mechanical stimuli (Hu et al., 2017). However, the cellular and molecular mechanisms underlying mechanical pain hypersensitivity remain enigmatic.

Recent studies demonstrate that epidermal cells work in concert with SSNs to transduce noxious and innocuous physical stimuli. For example, epidermal Merkel cells are mechanosensory cells that signal to sensory neurons to mediate touch transduction (Maksimovic et al., 2014; Hoffman et al., 2018). Similarly, keratinocytes are directly activated by noxious thermal and mechanical stimuli and release molecules that modulate nociceptor functions (Chung et al., 2004; Koizumi et al., 2004; Moqrich et al., 2005; Mandadi et al., 2009; Liu et al., 2019; Sadler et al., 2020). Furthermore, epidermal cells in invertebrates and vertebrates ensheath nociceptors in mesaxon-like structures (Cauna, 1973; Chalfie and Sulston, 1981; Han et al., 2012; Kim et al., 2012a; O’Brien et al., 2012; Jiang et al., 2019), and these sheaths may serve as sites of epidermis-nociceptor signaling (Yin et al., 2021). Indeed, epidermal ensheathment is required for normal responses to noxious mechanical stimuli in Drosophila (Jiang et al., 2019). However, whether epidermal cells are directly activated by noxious stimuli and modulate C4da neuronal activity has not been studied.

Here, we examined the capacity of Drosophila epidermal cells to drive nociceptor activation and modulate mechanical nociceptive responses. We found that stimulation of epidermal cells, but no other non-neuronal cell types in the larval body wall evokes activity in a variety of SSNs neurons and triggers nocifensive behavioral responses. Our in vitro and ex vivo calcium imaging experiments demonstrate that epidermal cells are intrinsically mechanosensitive. Using an unbiased genetic screen, we discovered a role for the store-operated calcium channel Orai, and its activator Stim in epidermal mechanotransduction and mechanical sensitization. Downstream of Stim/Orai activation, epidermal cells evoke nociceptor activation and mechanical hypersensitivity via epidermal vesicular release. Overall, we demonstrate that Drosophila epidermis-neuron signaling mediates both the acute detection of noxious mechanical stimuli and a form of prolonged mechanical hypersensitivity.

Results

Stimulation of epidermal cells evokes nocifensive behavior

To identify peripheral non-neuronal cell types that contribute to nociception, we conducted an optogenetic screen for light-evoked nocifensive behavior. First, as a benchmark for comparison we used the light-activated cation channel CsChrimson (Klapoetke et al., 2014) to optogenetically activate nociceptive C4da neurons. Consistent with prior reports (Hwang et al., 2007; Hu et al., 2017), C4da activation triggered nocifensive behaviors including c-bending and rolling in 100% of larvae (Fig. 1A, Fig. 1S1A, Movie S1). Next, we selectively expressed CsChrimson using GAL4 drivers in combination with elav-GAL80, which effectively siliences GAL4 expression in larval sensory neurons (Fig. 1S2), to target the six principle non-neuronal cell types within the larval body wall: epidermis, trachea, muscle, hemocytes, oenocytes, and glia (Fig. 1S3, Table S1). We then monitored light-evoked behavioral outputs associated with stimulation of each cell type. We found that optogenetic stimulation of epidermal cells, like C4da neurons, elicited nocifensive c-bending and/or rolling behaviors in 73% of larvae (Fig. 1A, 1S1B), without significantly altering nociceptor morphogenesis (Fig. 1S4). In contrast, stimulation of other body wall cell types elicited a variety of non-nociceptive behavior outputs: for example, muscle stimulation triggered hunching behavior followed by prolonged freezing, whereas glia stimulation reproducibly induced only hunching behavior (Fig. 1S1C-I) (Zimmermann et al., 2009). Thus, epidermal cells are the only non-neuronal body wall cell type that triggers robust nocifensive behavioral responses.

Figure 1.

To further validate the selective ability of body wall epidermal cells to elicit nocifensive behaviors, we examined eight other epidermal drivers in addition to R38F11-GAL4, which display no expression in sensory neurons and limited non-epidermal cell expression overall (Fig. 1S5). We found that optogenetic stimulation evoked nocifensive behaviors using each of the eight epidermal driver lines we tested: seven of the lines displayed rolling behavior while all eight displayed c-bending (Fig. 1B, Fig. 1S6). Although the previously described pan-epidermal A58-GAL4 driver (Galko and Krasnow, 2004) drove robust nocifensive rolling responses (Fig. 1B, 1S6), A58-GAL4 is expressed broadly in the larval CNS (Fig. 1S7) and stocchastically expressed in sensory neurons (Jiang et al 2014). In contrast, the remaining seven drivers including R38F11-GAL4 exhibited limited expression aside from epidermal cells, with no detectable expression in nociceptors, other larval SSNs, or peripheral glia, and highly restricted or undetectable expression in the CNS (Fig. 1S7). Further underscoring the connection between epidermal stimulation and nocifensive responses, the nocifensive behavioral response with these epidermal drivers correlated with the proportion of epidermal expression (Fig 1B).

We next used thermogenetic stimuluation with the warmth-activated TRP channel dTRPA1 (Hamada et al., 2008) as an independent method of probing nociceptive responses triggered by epidermal cell activation. On its own, the thermal stimulus (35° C) rarely induced rolling behavior in control larvae bearing UAS-TRPA1 alone. In contrast, we found that >75% of larvae expressing TRPA1 in all nociceptors exhibited rolling behavior in response to a thermal stimulus (Fig. 1C). Likewise, thermogenetic activation of epidermal cells induced robust rolling responses in >75% of larvae, and addition of GAL80 transgenes (tsh-GAL80 elav-GAL80) that silenced the sparse R38F11-GAL4 VNC expression (Fig. 1S5) had no effect on the rolling frequency (Fig. 1C, Fig. 1S3). Altogether, these results demonstrate that epidermal stimulation evokes nocifensive responses in Drosophila. Of note, prior studies demonstrated that sparse thermogenetic activation of nociceptors (<5 cells) yielded no significant increase in nocifensive rolling whereas activation of >10 cells elicited rolling responses in a majority of larvae (Robertson et al., 2013). Hence, epidermal stimulation likely engages numerous C4da neurons to elicit these behavioral responses.

In addition to C4da nociceptors, the epidermis is innervated by a variety of other SSNs including mechanosensory C3da and chordotonal (Cho) neurons and proprioceptive C1da neurons. Whereas direct stimulation of C4da nociceptors principally elicited nocifensive behavioral outputs, epidermal stimulation elicited an array of behaviors in addition to nocifensive responses, including freezing and hunching (Fig. 2A, 2B), behaviors associated with stimulation of C3da and Cho neurons (Zhang et al., 2013; Turner et al., 2016). These data suggest that epidermal cells may broadly modulate SSN activity in Drosophila.

Figure 2.

To examine whether different epidermis-evoked behaviors were associated with activation of distinct classes of SSNs, we compared epidermis-evoked and SSN-evoked behaviors. Stimulation of C4da, C3da and Cho neurons elicited distinct behavioral motifs: only C4da neurons elicited rolling behavior; stimulation of C3da and Cho neurons together elicited hunching, C-bending, and backing; stimulation of Cho neurons alone principally elicited hunching and freezing responses (Fig. 2A-2C, 2F). In contrast, optogenetic epidermal stimulation elicited all of these behaviors in a stereotyped sequence, with nocifensive behaviors (c-bending, rolling) preceding non-nociceptive behaviors (backing, freezing) (Fig. 2D, 2F, 2S1). We note that neither the behavioral motifs induced by epidermal or SSN stimulation nor the behavioral sequence induced by epidermal stimulation was recapitulated in effector-only controls (UAS-CsChrimson ATR+; Fig. 2S1E), demonstrating that the observed responses were driven by activation of the respective cell types.

We observed three striking differences in behavior evoked by stimulation of epidermal cells versus individual SSNs. First, although rapid, latency to rolling was significantly longer following epidermal stimulation compared to stimulation of C4da (Fig. 2F). Second, the duration of rolling, bending, and backing responses was significantly longer for epidermis versus SSN stimulation (Fig. 2G). Third, backing and freezing behaviors persisted beyond the duration of the light stimulus for epidermis but not SSN stimulation (Fig. 2H). In summary, we find that epidermal stimulation triggers more robust, varied and prolonged behaviors compared to responses from direct stimulation of discrete SSN subtypes.

Somatosensory neurons are activated by epidermal stimulation

We next asked whether epidermal stimulation activates larval SSNs including C4da, C3da, C1da, and Cho neurons. To test this possibility, we developed a semi-intact larval preparation in which we optogenetically stimulated epidermal cells while simultaneously monitoring calcium responses in axon terminals of SSNs (Fig. 3A). We found that epidermal stimulation triggered rapid and robust calcium transients in nociceptive C4da neurons, responses that were not observed in the absence of ATR or in effector-only controls (Fig. 3B). Epidermal stimulation likewise evoked calcium transients in mechanosensory C3da and Cho neurons, and in proprioceptive C1da neurons (Fig. 3C-3E, 3S1). Hence, epidermal stimulation can broadly modulate activity of larval SSNs.

Figure 3.

We next tested the requirement for SSN synaptic transmission in epidermis-evoked behaviors. We stimulated epidermal cells with CsChrimson while blocking SSN neurotransmitter release using tetanus toxin light chain (TnT) (Sweeney et al., 1995). We found that inhibiting C4da or C3da + Cho neurotransmission significantly reduced the frequency and duration of epidermal-evoked rolling and backing behaviors, respectively (Fig. 3E, 3F, 3S2). These data suggest that C4da and C3da/Cho neurons act downstream of epidermal cells to drive behaviors. We note that TnT expression in C4da neurons did not completely block epidermis-evoked nocifensive behaviors, and this likely reflects both incomplete C4da neuron silencing and epidermal activation of other SSNs that promote nociceptive outputs including C3da neurons, C2da neurons, and Cho neurons (Ohyama et al., 2015; Hu et al., 2017; Burgos et al., 2018). Further, silencing C4da or C3da/Cho neurons while stimulating epidermal cells led to an increase in the non-nocifensive behaviors hunching and freezing (Fig. 3E, 3F). These results, along with the observation that rolling behaviors predominate the early behavioral responses to epidermal stimulation (Fig. 2B), suggest that the nervous system prioritizes nocifensive behavioral outputs following epidermal stimulation. These data support a model in which epidermal cells and SSNs are functionally coupled.

Epidermal stimulation potentiates nociceptive neurons and behaviors

What is the physiological relevance of this functional coupling between epidermal cells and SSNs? To address this question, we compared calcium responses in C4da neurons to simultaneous epidermal and C4da stimulation or C4da stimulation alone. Simultaneous stimulation significantly enhanced the magnitude and duration of calcium responses in C4da axons (Fig. 4A-4D, 4S1). Based on this prolonged calcium response, we hypothesized that simultaneous epidermis and C4da neuron stimulation would yield enhanced nocifensive behavior output. To test this, we optogenetically stimulated C4da neurons and epidermal cells individually or in combination using low intensity CsChrimson activation and monitored larval behavior responses. In this stimulation paradigm, simultaneous epidermal cell and C4da neuron stimulation resulted in rolling in 100% of larvae whereas selective stimulation of C4da neurons or epidermal cells induced rolling in only 63% or 18% of larvae, respectively (Fig. 4E, 4F). Furthermore, simultaneous stimulation elicited a significantly higher number of rolls among responders than stimulation of nociceptors or epidermal cells alone (26.9 rolls for C4da + Epi, 4.9 for C4da, and 5.3 for Epi stimulation; Fig 4G, 4H). Likewise, simultaneous stimulation significantly reduced the latency to the first roll (Fig. 4I) and increased the duration of rolling behaviors (Fig. 4J). We next tested whether this functional coupling extends to mechanical stimuli. We simultaneously presented larvae with a noxious mechanical stimulus and a low intensity optogenetic epidermal stimulus that was insufficient to trigger rolling on its own (0% response rate, n = 200). This concurrent epidermal stimulation significantly increased touch-evoked nocifensive responses, yielding a 91% or 49% increase in rolling responses to 20 mN or 50 mN Von Frey stimulus, respectively (Fig. 4K). We next probed the kinetics of this epidermis-induced mechanical sensitization.

Figure 4.

When Drosophila larvae are presented with two nociceptive mechanical stimuli in succession, they exhibit enhanced behavioral responses to the second stimulus (Hu et al., 2017). We hypothesized that selective epidermal stimulation would sensitize larvae to subsequent nociceptive mechanical stimuli. To test this hypothesis, larvae expressing the warmth-activated calcium-permeable channel dTRPA1 in epidermal cells were presented with a thermal stimulus, 32° C to activate dTRPA1, followed by a 40 mN mechanical stimulus 10 sec later (Fig. 4L). Indeed, we found that dTRPA1-mediated epidermal stimulation significantly sensitized larvae to a subsequent mechanical stimulus, increasing the roll probability more than two-fold. In contrast, dTRPA1-mediated stimulation of C4da neurons did not induce mechanical sensitization, and we confirmed this result with two independent C4da neuron drivers (Fig. 4L). Thus, activation of epidermal cells but not C4da nociceptors alone induces prolonged sensitization to noxious mechanical stimuli. We next assessed the duration of sensitization following transient epidermal activation. Thermogenetic epidermal stimulation yielded persistent sensitization that recovered over a timescale of minutes (τ = 337 sec, Fig. 4M, 4N). The magnitude and duration of mechanical sensitization by thermogenetic epidermal stimulation was remarkably similar to the sensitization evoked by a prior mechanical stimulus (63% roll probability in response to a second stimulus, τ = 334 sec, Fig. 4N, 4S1B). Altogether our data support a model whereby epidermal cells are mechanosensitive cells that signal to SSNs to drive acute nocifensive behaviors and prolong mechanical sensitization.

Epidermal cells are intrinsically mechanosensitive

Prior studies have shown that vertebrate epidermal cells directly respond to mechanical stimuli (Koizumi et al., 2004; Haeberle et al., 2008; Tsutsumi et al., 2009; Ranade et al., 2014; Woo et al., 2014; Moehring et al., 2018). Therefore, we next assessed whether Drosophila epidermal cells are intrinsically mechanosensitive. We developed a protocol to acutely dissociate epidermal cells and measure the responses of individual GCaMP6s-expressing epidermal cells to mechanical stimuli (Fig. 5A). We found that radial stretch elicits calcium responses in epidermal cells in a dose-dependent manner. For example, a low 0.5% stretch activated 18% of cells and a subsequent 1% stretch recruited an additional 10% of stretch-responding cells (Fig. 5B-5D). Overall, 51% of epidermal cells displayed stretch sensitivity (Fig. 5C, 5D). We also found that 43% of epidermal cells responded to hypoosmotic challenge and 35% responded to laminar flow; 19% of epidermal cells responded to both hypoosmotic challenge and laminar flow (Fig. 5S1). Given that dissociated epidermal cells were intrinsically mechanosensitive, we next assessed mechanically evoked responses in a semi-intact body wall preparation (Fig. 5E). We found that 50% of epidermal cells exhibited a robust calcium transient in response to a 25 μm membrane displacement using a glass probe (Fig. 5E-5G, 5S1G). Altogether, these results indicate that Drosophila larval epidermal cells are intrinsically mechanosensitive.

Figure 5.

Mechanically evoked epidermal responses rely on store-operated calcium entry

Our studies demonstrate that, like vertebrate keratinocytes, Drosophila epidermal cells exhibit mechanically evoked calcium transients. What is the mechanism of mechanotransduction in these cells? RNA-seq analysis of acutely dissociated epidermal cells revealed expression of more than 20 cation channels, including the mechanosensitive ion channels Piezo, TMEM63, and TMCO (Fig. 6S1). We assessed the epidermal requirements of these channels in mechanical nociception using available RNAi transgenes (Fig. 6A). Our behavioral screen identified one channel, Orai, the sole Drosophila pore-forming subunit of the Ca²⁺ release-activated Ca²⁺ (CRAC) channel (Feske et al., 2006), that blocked mechanically-evoked nociceptive sensitization without impacting behavioral responses to the first stimulus (Fig. 6A, 6B, 6S2A) or altering nociceptor morphogenesis (Fig. 1S4). Interestingly, our screen uncovered an epidermal role for Task6, an orthologue of stretch-sensitive 2-pore potassium channels (Fink et al., 1996), in mechanonociception, as Task6 RNAi increased nocifensive rolling responses to the initial mechanical stimulus (Fig. 6S2B). Finally, although our RNAi studies did not reveal epidermal requirement for other known mechanosensitive cation channels in mechanonociceptive behaviors, it is possible that multiple channels function redundantly, or that RNAi knockdown was incomplete.

Figure 6.

To gain insight into mechanically evoked nociceptive sensitization, we focused on probing the role of Orai in epidermal mechanosensory responses. We first asked whether Orai is functional in Drosophila epidermal cells. Orai is a store-operated calcium (SOC) channel that is activated by the endoplasmic reticulum (ER)-calcium sensitive molecule, Stim, upon calcium release from ER calcium stores. Thapsigargin (TG) induces calcium release from intracellular stores and thus triggers activation of Orai channels. Indeed, Drosophila epidermal cells displayed TG-induced calcium release in the absence of extracellular calcium, followed by calcium influx upon re-addition of extracellular calcium (Fig. 6C). Calcium influx was significantly inhibited by the addition of low nanomolar lanthanum, consistent with the high sensitivity of Orai channels to lanthanides (Fig. 6S2C). This characteristic store operated calcium entry (SOCE) response was significantly reduced by epidermis-specific Stim or Orai RNAi knockdown (Fig. 6S2D-F). Consistent with a key role for SOCE in mechanotransduction, we found that radial stretch induced both calcium release from intracellular stores in the absense of extracellular calcium and influx upon calcium re-addition. While both store release and calcium influx constitute the calcium response to stretch, in 69% of cells, calcium due to store release exceeded that of calcium re-entry (Fig. 6E). Consistent with this observation, depletion of intracellular stores and inhibition of calcium re-entry reduced the number of stretch sensitve cells by 61% (stretch non-responsive cells in control = 49% vs. store depleted = 80%) and 30% (stretch non-responsive cells in control = 49% vs. La³⁺ = 64%; Fig. 6F-G), respectively. Given that Stim and Orai mediate SOCE in epidermal cells and that stretch evokes SOCE, we investigated requirements for epidermal Stim and Orai in mechanically evoked calcium responses. RNAi knockdown of either Stim or Orai significantly reduced the fraction of stretch-responsive epidermal cells (RNAi control = 48%, Stim RNAi = 22%, Orai RNAi = 24%; Fig. 6H-I), with Stim or Orai RNAi preferentially attenuating stretch evoked responses to larger magnitude stretch stimuli. We also found that human keratinocytes display dose-dependent stretch evoked calcium responses, though they respond to higher magnitudes of stretch than Drosophila epidermal cells (Fig. 6J). Like Drosophila, both store release and calcium influx contribute to stretch evoked calcium responses in human keratinocytes (Fig. 6K).

Two hallmarks of Stim and Orai-mediated SOCE are steep inward rectification, with larger currents at hyperpolarizing potentials, and highly cooperative Orai activation by Stim (Hoover and Lewis, 2011). Since Stim and Orai mediate mechanical responses of epidermal cells in vitro, we predicted that increasing the calcium driving force through Orai activity by either hyperpolarizing epidermal cells or by activating additional Orai channels via Stim overexpression would enhance behavioral responses to mechanical stimuli. Indeed, we found that hyperpolarizing epidermal cells with the light-activated anion channelrhodopsin GtACR1 (Mohammad et al., 2017) increased behavioral responses to mechanical stimuli (Fig. 6L). In addition, overexpressing Stim in epidermal cells significantly enhanced nocifensive behavioral responses to mechanical stimuli (Fig. 6M). Altogether, these results demonstrate that mechanically evoked responses of epidermal cells and the resulting nocifensive behavior outputs require store-operated calcium entry.

How might mechanically evoked calcium entry in epidermal cells drive nociceptor activation and behavior? Stim/Orai-mediated calcium entry contributes to exocytosis in a variety of cell types, including neurons and immune cells (Pores-Fernando and Zweifach, 2009; Ashmole et al., 2012; Maneshi et al., 2020; Chanaday et al., 2021; Ramesh et al., 2021). Therefore, we investigated the contribution of epidermal exocytosis in nociceptive sensitization with the temperature-sensitive dynamin mutant shibire^ts (shi^ts) to inducibly block vesicle recycling, as this treatment rapidly and potently blocks neurotransmitter release (Koenig et al., 1983) and we found that acute epidermal dynamin inactivation using UAS-shi^tshad no discernable effect on nociceptor morphogenesis (Fig. 1S4). In this paradigm, larvae expressing shi^ts in epidermal cells, but not control larvae, exhibited significant attenuation of mechanically evoked nociceptive sensitization following pre-incubation at the non-permissive temperature (Fig. 6N). In contrast, both genotypes exhibited comparable responses to a mechanical stimulus at the permissive temperature (25° C) and to the first mechanical stimulus following pre-incubation at the non-permissive temperature (30° C). Taken together, these results are consistent with a model in which mechanical stimuli induce calcium influx and vesicular release from epidermal cells, which in turn activates nociceptors to induce acute nocifensive behaviors and prolonged sensitization (Fig. 6O). Although our RNA-seq analysis of epidermal cells did not reveal expression of neurotransmitter biosynthesis genes, epidermal cells express a large repertoire of genes involved in vesicular release as well as several neuropeptide genes, providing an entrypoint to defining the molecules involved in epidermis-SSN communication.

Discussion

In this study, we have shown an essential role for Drosophila epidermal cells in escape responses to noxious mechanical stimuli. Activation of epidermal cells acutely activates SSNs to induce an array of behavioral outputs and mechanical sensitization. This epidermal potentiation persists for minutes to promote a prolonged, but reversible, mechanical hypersensitivity that may protect from further insult. This is distinct from previously described forms of neuropathic thermal and mechanical hypersensitivity in Drosophila which are induced by tissue damage and chemotherapuetic agents, respectively, emerge on a timescale of hours, and are long-lasting (Babcock et al., 2009; Boiko et al., 2017; Khuong et al., 2019). In the mammalian somatosensory system, a variety of inflammatory mediators have been shown to activate TRPA1 in neurons to promote mechanical hypersensitivity (Bautista et al., 2006); however, the molecular force transducers that mediate mechanical pain are unknown. In contrast, in the Drosophila somatosensory system, Ppk1/Ppk26, Piezo, and Trpa1 are key transducers of mechanonociception (Zhong et al., 2010; Kim et al., 2012b; Gorczyca et al., 2014; Guo et al., 2014; Mauthner et al., 2014). Prolonged sensitization to noxious mechanical stimuli plays an important protective role in an organism’s survival; yet the mechanisms of mechanical sensitization of Drosophila nociceptors were unknown.

We demonstrate a new role for SOCE signaling in both Drosophila and human epidermal cell mechanotransduction. While short-term sensitization is beneficial to survival, a key hallmark of pathological pain is prolonged and persistent mechanical hypersensitivity; whether deregulation of this mechanism of epidermis-evoked short-term sensitization contributes to pathological pain remains to be determined. Overall, we identified a mechanism that does not impact acute nociception but selectively regulates mechanical sensitization. These findings highlight Stim/Orai signaling as a new avenue for understanding mechanical pain.

This work has opened several new directions for future studies. First, how does radial and osmotic stretch lead to the activation of store-operated calcium signaling? Although Orai has not previously been shown to be mechanosensitive, our studies revealed a requirement for Orai and its activator Stim in mechanically-evoked calcium flux in Drosophila epidermal cells. We also showed that radial stretch of human keratinocytes triggered calcium release from stores and SOCE; our previous studies showed that Stim and Orai are required for SOCE in human keratinocytes (Wilson et al., 2013). These data in combination with previous findings that showed mechanical stimulation of human mesenchymal stem cells also triggers SOCE (Kim et al., 2015) suggests that Stim/Orai signaling may represent a conserved pathway for mechanotransduction in non-neuronal cells.

Second, how is Stim/Orai function linked to mechanotransduction? Stim/Orai signaling is activated downstream of G-protein coupled receptors (GPCRs) and receptor tyrosine kinases (RTKs) through phospholipase C. Studies have shown that a number GPCRs are mechanosensitive (Chachisvilis et al., 2006; Grosmaitre et al., 2007; Mederos y Schnitzler et al., 2008; Connelly et al., 2015; Xu et al., 2018), though this has not been studied in epidermal cells. Alternatively, plasma membrane deformation has been shown to induce ER-plasma membrane junctions (Venturini et al., 2020; Aoki et al., 2021), where Stim and Orai clusters accumulate and interact to drive calcium influx (Luik et al., 2008).

Third, how does mechanically-induced singaling in epidermal cells lead to modulation of SSNs? Our data support a model whereby epidermal cells and multiple classes of SSNs are functionally coupled. Epidermal stimulation modulates activity of nociceptive C4da neurons, mechanosensory C3da and Cho neurons, and proprioceptive C1da neurons, and the output of neuronal activity is required for epidermis-evoked behaviors. We demonstrated a requirement for dynamin-dependent vesicle release from epidermal cells in mechanical sensitization, providing a potential link between Stim/Orai signaling in epidermal cells and downstream neuronal activity. However, the mediators that are released by epidermal cells and the signaling molecules in the nociceptors remain unknown. Furthermore, whether different subtypes of SSNs are coupled to epidermal cells by distinct mechanisms remains to be determined. At least in the case of Cho neurons which are wrapped by ensheathing glial cells and scolopale cells, signaling from epidermal cells likely involves at least one additional cell type.

Epidermal cells ensheath peripheral arbors of some SSNs, including Drosophila nociceptive C4da neurons and, to a lesser extent, mechanosensory C3da neurons (Jiang et al., 2019). Hence, epidermal sheaths could facilitate transduction of epidermal signals that modulate nociceptor function. Consistent with this possibility, blocking ensheathment attenuates Drosophila larval responses to noxious mechanical stimuli (Jiang et al., 2019) and likewise impairs function of some C. elegans mechanosensory neurons (Chen and Chalfie, 2014). However, our finding that epidermal stimulation evokes calcium responses from SSNs that are not ensheathed by epidermal cells (C1da, Cho neurons) argues that epidermal sheaths are unlikely to play an essential function in epidermis-SSN functional copuling. Instead, ensheathment may facilitate nociceptor activation by increasing the efficiency of vesicular exchange or, alternatively, may modulate nociceptor activity through through enhanced ionic coupling to epidermal cells.

Which epidermal-derived molecules might modulate neuronal activity? There are several mechanisms by which mammalian epidermal cells activate SSNs. Vesicular release of norepinephrine from mouse epidermal Merkel cells is required for sustained touch-evoked firing of mechanosensory neurons (Hoffman et al., 2018). Additionally, mechanical stimuli trigger ATP release from mouse keratinocytes that activates nociceptors via purinergic (P2X4) receptors (Koizumi et al., 2004; Tsutsumi et al., 2009; Moehring et al., 2018). Finally, Stim/Orai-dependent SOCE mediates the release of the cytokine thymic stromal lymphopoietin (TSLP) from keratinocytes that directly activates a subset of TRPA1-expressing SSNs to induce itch (Wilson et al., 2013). Similar to these mammalian models, UV-damage has been shown to induce the release of the cytokine Eiger to promote nociceptor sensitization (Babcock et al., 2009); though this occurred on a slower timescale than the epidermal-evoked mechanical sensitization we describe here (8 hrs vs. ∼ 10 sec, Fig. 4L). Likewise, epidermal platelet-derived growth factor (PDGF) ligands regulate mechanonociceptive responses in Drosophila (Lopez-Bellido et al., 2019) and intrathecal delivery of PDGF or the closely related growth factor EGFR yields mechanical hypersensitivity in rats (Masuda et al., 2009; Puig et al., 2020), but it remains to be determined whether growth factor signaling can yield rapid sensitization. Hence, future studies will address which neurotransmitters, neuropeptides, or inflammatory mediators underly epidermal cell-mediated mechanical sensitization.

Our data support a model whereby epidermal cells and multiple classes of SSNs are functionally coupled. Future studies will address which neurotransmitters, neuropeptides, or inflammatory mediators underly epidermal cell-mediated mechanical sensitization. An additional key next step is understanding whether the neuronal plasticity underlying mechanical sensitization results from the direct modulation of mechanosensitive channels or rapid insertion of new mechanosensitive channels into the plasma membrane, or from changes in the signaling pathways or channels that regulate neuronal excitability. Overall, we performed an unbiased genetic screen that for the first time establishes a key role for mechanically-evoked Stim/Orai calcium signaling in epidermal cells that drive nociceptor modulation and mechanical hypersensitivity.

Material and Methods

Drosophila strains

Flies were maintained on standard cornmeal-molasses-agar media and reared at 25° C under 12 h alternating light-dark cycles. For all experiments involving optogenetic manipulations, larvae were raised in the constant dark at 25 °C on Nutri-Fly Instant Food (Genesee Scientific #66-117), supplemented with 1 mM all-trans retinal (ATR; Sigma #R2500). See Table S1 for a complete list of alleles used in this study. Experimental genotypes are listed in figure legends.

Behavior analysis

Optogenetic behavior screen

Individual larvae were rinsed in ddH2O, transferred to an agarose substrate (1% agarose, 100 mM dish) in a darkened arena, and habituated for 30 sec. Larvae were stimulated with a top-mounted 488 nM LED illuminator (PE-300, CoolLED) and images were captured with a sCMOS camera (Orca Flash 3.0, Hamamatsu) at frame acquisition rate of 20 fps and behaviors were scored before, during and after optogenetic stimulation.

High resolution video tracking of optogenetic-gated larval behavior

Following 5 min of light deprivation including 15 sec of habituation in the behavioral arena, larvae were tracked before, during and after optical stimulus (10 sec each, 30 sec total) (Fig. 2A). For these studies we modified our stimulation paradigm in two key ways: to avoid potential contributions of nociceptor light evoked responses (Xiang et al., 2010), we stimulated larvae using yellow-shifted light; and to facilitate kinetic analysis of behavior outputs, we used an automated shutter. Larvae were stimulated with a top-mounted 585 nm LED illuminator (SPECTRA X, Lumencor) equipped with a filter (FF01 585/40-25, Semrock), and images were captured with a sCMOS camera (Zyla4.2, Andor) at a frame rate of 20Hz. Larvae were constantly illuminated with an infrared (940 nm) light source (LDR2-132IR2-940-LA, CSS) for visualization. Larvae were fed (ATR+) or vehicle alone (ATR-) as indicated.

Thermogenetic behavior assays

Larvae for thermogenetic assays were reared at room temperature (20° C) to limit TRPA1 activation during development. Third instar larvae were isolated from their food, washed in distilled water, and recovered to damp agar plates for several min, and transferred individually to a Peltier plate held at 25° C or 35° C. Behavior responses were recorded under infrared light with a computer-controlled GigE camera (FLIR) at an acquisition rate of 20 fps for 20 sec. Responses were analyzed post-hoc blind to genotype and were plotted as the proportion of larvae that exhibited at least one complete nocifensive roll during stimulus application.

Mechanonociception assays

Third instar larvae were isolated from their food, washed in distilled water, and placed on a scored 35 mm petri dish with a thin film of water such that larvae stayed moist but did not float. Larvae were stimulated dorsally between segments A4 and A7 with calibrated Von Frey filaments that delivered the indicated force upon buckling, and nocifensive rolling responses were scored during the 10 sec following stimulus removal. For assays involving multiple stimuli, larvae were stimulated individually, allowed to freely locomote in the arena for up to 1 min (for longer recoveries larvae were recovered onto 2% agar to prevent desiccation), and subsequently presented with the second stimulus. For assays involving thermal and mechanical stimuli, larvae were individually transferred to a pre-warmed Peltier plate containing a thin layer of water, incubated for the indicated time, and transferred to the behavior arena (or a 2% agar plate for recoveries > 1 min) with a paint brush for subsequent mechanical stimulation. All assays were conducted in ambient light except for experiments with GtACR (Fig. S72), which were conducted under broad-spectrum (500-700 nm) LED illumination (CoolLED PE-300, green). Our illumination setup for these experiments provided limited working distance, therefore larvae were restrained with forceps and given only a single stimulus.

Video annotations

Videos of individual larvae responding to light stimuli were scored on a frame-by-frame basis using the annotation software BORIS (Friard and Gamba, 2016). Behaviors scored, along with descriptions of the criteria for each behavior, are detailed in Table S2. Video analysts were blind to the genotype and treatment during scoring. Scoring on a training set was compared across all analysts to calibrate, and any behaviors for which the primary analyst was uncertain were reviewed by an additional analyst. Additionally, 10% of videos were scored independently by two analysts and there was at least 80% concordance in behaviors annotated in these comparisons.

Microscopy

Calcium imaging: ventral nerve cords

Third-instar larvae were dissected along the dorsal midline and pinned on a sylgard-coated dish (Silpot 184, Dow Corning Toray). The internal organs except for neural tissues were removed. Larvae were bathed in HL3.1 (Feng et al, 2004) modified to remove calcium (Table S3) to minimize larval movement. The ventral nerve cord was imaged using an Olympus BX51WI microscope, equipped with a spinning-disk confocal unit Yokogawa CSU10 (Yokogawa) and an EM-CCD digital camera (Evolve, Photometrics). For activation of epidermal cells with the light gated CsChrimson, red light was delivered by a pE-300 (CoolLED) equipped with a filter (ET645/30x, Chroma). Obtained images were analyzed using Metamorph (https://www.moleculardevices.com/systems/metamorph-research-imaging) and ImageJ (Schneider et al., 2012). Baseline fluorescence was calculated as the mean fluorescence intensity of an ROI over the ten frames prior to light stimulus delivery. The trapezoidal method was used to calculate area under the curve, utilizing the trapz function of MATLAB. Data points from the onset of stimulation to the end of stimulation were used for the calculation.

Calcium imaging: fillet preparations

Third-instar larvae were dissected along the ventral midline and pinned on sylgard (Dow Corning) dishes with the internal surface facing towards the microscope. All internal organs, including the central nervous system, were removed. Larvae were bathed in calcium-containing HL3.1 (Feng et al., 2004) (Table S3) except where indicated and images of the dorsal midline between abdominal segments A2 and A4 were captured with a Zeiss Axio Zoom V16 microscope. Captured images were analyzed using ImageJ (Schneider et al., 2012). Mechanical stimulus: fillets were poked with a tapered borosilicate capillary with a rounded tip, using a micromanipulator to induce a deflection of 25 µm. The decay time constant was calculated by fitting the data points from the peak response to the end of the experiment into an exponential curve f(x) = a*exp(b*x) using MATLAB with R²>0.9 used as a threshold for reliable fitting.

Calcium imaging: dissociated epidermal cells

Six to eight larval fillets were dissociated in 400 µL of 50% Saline (modified Ringer’s recipe) / 50% Schneider’s media with 200 U/mL collagenase type I (Fisher 17-100-017), with mixing at 1000 RPM at 33°C for 16 min, with trituration every 8 min. Undigested fillets were removed and the remaining suspension was spun at 500 g for 3 min, followed by aspiration of the supernatant down to a 10 µL cell suspension. Cells were resuspended in 30 µL fresh PBS / Schneider’s solution and plated onto poly-D-lysine (1 mg/ml, Sigma P7886) coated No. 1 coverslips, with 10 µL cell solution per coverslip. Cells were cultured at least 30 min and up to 2 hr at 25° C prior to imaging. Cells were imaged using a a 10x objective at a frame rate of 0.33 Hz. Solutions are indicated in figure legends (see Table S3 for recipes). Obtained images were analyzed using MetaFluor and Python and baseline fluorescence was calculated as the mean fluorescence intensity of an ROI over 5 frames prior to stimulus delivery. For stretch stimulation, circular membranes were cut with an arch punch from sheets of glossy silicone of 0.01–0.02 inch thickness (Specialty Manufacturing, Inc.) and coated with 1 mg/ml poly-D-lysine for 1 h before plating cells. Membranes were mounted onto the StageFlexer system and vacuum pressure was applied through the FX-3000 system (Flexcell). Calibrations were performed using fluorescent beads attached to the membranes, and images were taken before and during a static stretch. To stimulate cells, a 2 sec square wave of vacuum pressure was applied. Cells were imaged with an Olympus BX61WI upright microscope. For store-operated calcium entry measurements and osmotic stimulation, cells were imaged using a Zeiss Observer inverted microscope and solutions were perfused using the Automate Scientific ValveLink 8.2 perfusion system. At the end of each imaging session, 1uM ionomycin was perfused and only cells that showed a calcium response, as defined by a 10% increase from baseline fluorescence, were used in analysis. Flow, osmotic and radial stretch responders were defined by a 5% increase from baseline fluorescence.

Calcium imaging: human keratinocytes

Immortalized human keratinocytes (HaCaT) cells were plated on silicone membranes one day prior to stretch experiments. Prior to the radial stretch experiments, cells were loaded with 1 μm Fura-2AM supplemented with 0.01% Pluronic F-127 (w/v, Life Technologies) in a physiological Ringer’s solution containing the following (in mm): 140 NaCl, 5 KCl, 10 HEPES, 2 CaCl2, 2 MgCl2, and 10 d-(+)-glucose, pH 7.4. Acquired images were displayed as the ratio of 340 nm/380 nm. Cells that had a response 10 standard deviations above baseline to ionomycin were included in the analysis and stretch responses were defined by a 15% increase in Fura-2 340/380 ratio.

Confocal Microscopy

For peripheral imaging of cellular morphology, live single larvae were mounted in 90% glycerol under a coverslip and imaged on a Leica SP5 confocal microscope using a 40x 1.25 NA lens. To image the larval CNS, larvae were dissected on sylgard plates, briefly fixed in 4% paraformaldehyde (PFA) in PBS for 15 min at room temperature, washed 3 x 5 min in PBS, and mounted for imaging.

RNA-Seq analysis of epidermal cells

RNA isolation for RNA-Seq

Larvae with cytoplasmic GFP expressed in different epidermal subsets were microdissected and dissociated in collagenase type I (Fisher 17-100-017) into single cell suspensions, largely as previously described (Williams et al., 2016), with the addition of 1% BSA to the dissociation mix. After dissociation, cells were transferred to a new 35 mm petri dish with 1 mL 50% Schneider’s media, 50% PBS supplemented with 1% BSA. Under a fluorescent stereoscope, individual fluorescent cells were manually aspirated with a glass pipette into PBS with 0.5% BSA, and then serially transferred until isolated without any additional cellular debris present. Ten cells per sample were aspirated together, transferred to a mini-well containing 3ul lysis solution (0.2 % Triton X-100 in water with 2 U / µL RNAse Inhibitor), lysed by pipetting up and down several times, transferred to a microtube, and stored at -80° C. For the picked cells, 2.3 µL of lysis solution was used as input for library preparation.

RNA-Seq library preparation

RNA-Seq libraries were prepared from the picked cells following the Smart-Seq2 protocol for full length transcriptomes (Picelli et al., 2014). To minimize batch effects, primers, enzymes, and buffers were all used from the same lots for all libraries. Libraries were multiplexed, pooled, and purified using AMPure XP beads, quality was checked on an Agilent TapeStation, and libraries were sequenced as 51-bp single end reads on a HiSeq4000 at the UCSF Center for Advanced Technology.

RNA-Seq data analysis

Reads were demultiplexed with CASAVA (Illumina) and read quality was assessed using FastQC (https://www.bioinformatics.babraham.ac.uk/) and MultiQC (Ewels et al., 2016). Reads containing adapters were removed using Cutadapt version 2.4 (Martin, 2011) and reads were mapped to the D. melanogaster transcriptome, FlyBase genome release 6.29, using Kallisto version 0.46.0 (Bray et al., 2016) with default parameters. AA samples were removed from further analysis for poor quality, including low read depth (< 500,000 reads), and low mapping rates (< 80%). Raw sequencing reads and gene expression estimates are available in the NCBI Sequence Read Archive (SRA) and in the Gene Expression Omnibus (GEO) under accession number SUB11489996.

Statistical analysis

For each experimental assay control populations were sampled to estimate appropriate sample numbers to allow detection of ∼33% differences in means with 80% power over a 95% confidence interval. Details of statistical tests including treatment groups, sample numbers, statistical tests, p-values and q-values are provided in the experimental supplement.

Acknowledgements

This work was supported by grants from the National Institutes of Health to J.Z.P. (NINDS R01 NS076614), DB (NICHD K99 HD086271), CW (5F31NS106775), and the MBL (R25NS063307); a grant from the National Science Foundation to S.S.M (NSF GRFP DGE1752814); a grant from the Weill Neurohub to J.Z.P and D.B.; a grant from the Scan Design Foundation, a JSPS long-term fellowship and startup funds from UW (J.Z.P); MEXT Grants-in-Aid for Scientific Research (KAKENHI 16H06456), JSPS (KAKENHI 16H02504), WPI-IRCN, AMED-CREST (JP22gm310010), and JST-CREST to KE; and a fellowship from the Grass Foundation (C.E.E.). Fly Stocks obtained from the Bloomington Drosophila Stock Center (NIH P40OD018537) were used in this study. We thank Jessica Huang, Jordan Martel, and David Shen for assistance with video tracking; Peter Soba for helpful discussions.

Author Contributions

Conception and design: Optogenetic behavior analysis, K.E., J.Z.P, C.R.W, J.Y.; larval behavior assays, J.Z.P, C.R.W, and J.Y.; in vivo calcium imaging, K.E., J.Z.P, C.R.W, J.Y.; ex vivo calcium imaging, D.B., S.S.M., and J.Z.P

Acquisition of data: Optogenetic behavior screen, C.J.A, C.E.E., S.M., and J.Y.; Optogenetic behavior assays, J.Y., C.H., M.M., K.I., and C.R.W.; mechanonociception assays, C.R.W, and J.Z.P; thermogenetic behavior assays, J.Z.P.; RNA-sequencing, J.Z.P. and C.R.W.

Analysis and Interpretation of data: larval behavior assays, J.Z.P, C.R.W, J.Y.; in vivo calcium imaging, K.E., J.Z.P, C.R.W, J.Y.; ex vivo calcium imaging, D.B. and S.S.M.; transcriptomic data, C.H., C.R.W., and J.Z.P.

Drafting the article: D.B., J.Z.P, C.R.W, S.S.M., and J.Y.

Competing Interests

DMB is on the scientific advisory board of Escient Pharmaceuticals. The remaining authors declare no conflicts of interest.

Details of Statistical Analysis

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