We first obtained whole-cell patch clamp recordings from acutely dissociated nematocytes to investigate nematocyte signal transduction. Using intracellular cesium (Cs⁺) to block potassium (K⁺) currents revealed a voltage-gated inward current that was activated by positive or depolarized membrane voltages (I_CaV, Figure 1C). In response to sustained positive voltage, voltage-activated ion channels enter a non-conducting, inactivated state and cannot be activated until returned to a resting state by negative membrane voltage. This property generally serves to limit responses to repetitive or prolonged stimulation, similar to receptor desensitization. Remarkably, I_CaV began to inactivate at voltages more negative than we could technically measure, thus demonstrating an unusual voltage sensitivity of this conductance (Figure 1C,D). To determine whether these properties were specific to nematocytes, we used a transgenic sea anemone with fluorescently labeled neurons to facilitate direct comparison between these excitable cell types (Nakanishi et al., 2012; Figure 1C). Neuronal voltage-gated currents had a lower threshold for activation and exhibited much weaker voltage-dependent inactivation (Figure 1C,D, Figure 1—figure supplement 1A–D), similar to currents found in neurons of other animals (Hille, 2001), indicating that nematocytes exhibit unusual voltage-dependent properties. Ion substitution and pore blocker experiments confirmed I_CaV is a Ca²⁺-sensitive current (Figure 1E,F), consistent with the contribution of extracellular Ca²⁺ to chemo-tactile-induced discharge (Watson and Hessinger, 1994a; Gitter et al., 1994; Figure 1G). Increased concentrations of extracellular Ca²⁺ did not affect inactivation of I_CaV (Figure 1—figure supplement 1E), suggesting the enhanced voltage-dependent inactivation is intrinsic to the channel complex. This observation is important because it suggests I_CaV renders nematocytes completely inactivated at typical resting membrane voltages and thus cells could not be stimulated from resting state.

To identify the ion channel mediating I_CaV, we generated a tentacle-specific transcriptome and aligned reads from nematocyte-enriched cells (Sunagar et al., 2018). This strategy allowed us to search for differentially expressed transcripts that might encode Ca_V channel subunits (pore-forming α and auxiliary β and α2δ subunits). The orthologue of cacnb2, a β subunit of Ca_V channels, was the highest expressed Ca_V transcript in nematocyte-enriched cells, with levels 14-fold higher than other cells in the sea anemone (Figure 2A). β subunits can modulate voltage-dependence and trafficking in diverse ways depending on their splice isoform, interacting subunits, and cellular context (Buraei and Yang, 2010). Importantly, β subunits only interact with α subunits of high voltage-activated (HVA) calcium channels (Perez-Reyes, 2003). In agreement with robust β subunit expression, we found significant enrichment for cacna1a, the pore-forming subunit of HVA Ca_V2.1, and high expression of cacna2d1 (Figure 2—figure supplement 1A,B). These observations are consistent with a previous report demonstrating specific expression of cacna1a in nematocytes of sea anemone tentacles and expression of β subunits in nematocytes from jellyfish (Bouchard et al., 2006; Moran and Zakon, 2014; Bouchard and Anderson, 2014). Expression of cacna1h, which does not interact with auxiliary subunits (Buraei and Yang, 2010), was also observed, albeit at lower levels and across all cells (Figure 2—figure supplement 1A). Thus, it remains possible that voltage-gated currents in nematocytes are not carried exclusively by one Ca_V subtype.

Heterologous expression of Nematostella Ca_V (nCa_V: cacna1a, cacnb2, and cacna2d1) produced voltage-gated currents with an apparent activation threshold nearly identical to the Ca_V complex made from respective mammalian orthologues (mCa_V, Figure 2B, Figure 2—figure supplement 1C,D). Both channels had similar activation kinetics, but fast inactivation was significantly pronounced in nCa_V, resembling native I_CaV (Figure 2E, Figure 2—figure supplement 1E). Importantly, nCa_V voltage-dependent inactivation was greatly enhanced compared with mCa_V, regardless of the charge carrier (Figure 2B, Figure 2—figure supplement 1F,G). Similar to I_CaV, nCa_V exhibited unusually-sensitive voltage-dependence and began to inactivate at voltages more negative than we could measure with an estimated midpoint inactivation voltage (V_i1/2)~80 mV more negative than mCa_V (Figure 2B). Even with a holding potential of −70 mV, nCa_V exhibited slow inactivation resulting in a drastic decrease in responses to depolarizing stimuli over time (Figure 2C). This slow inactivation was largely prevented by adjusting the holding potential to −110 mV, suggesting inactivation occurs when channels are in a closed-state at potentials near or more negative than typical resting membrane potential (Figure 2—figure supplement 1H). Importantly, nCa_V rapidly recovered from inactivation, demonstrating that channels could be reset for subsequent activation following brief exposure to negative voltage (Figure 2D). These distinctive features closely match the unique properties of native I_CaV, suggesting nCa_V forms the predominant Ca_V channel in nematocytes.

To determine the molecular basis for nCa_V inactivation, we analyzed chimeric Ca_V complexes containing specific α, β, and α2δ1 subunits from nCa_V or mCa_V orthologues. Using a holding potential of −110 mV to compare voltage-dependent inactivation, we found that only transfer of the β subunit significantly affected voltage-dependent inactivation, while α or α2δ1 subunits produced minimal effects on voltage-dependent activation, inactivation, or kinetics (Figure 2E, Figure 2—figure supplement 1C–E). Indeed, other β subunits can induce significant hyperpolarized shifts in inactivation of HVA Ca_V channels (Yasuda et al., 2004). In this case, the mCa_V β subunit drastically shifted nCa_V inactivation by ~56 mV in the positive direction, prevented complete inactivation, and produced slower fast inactivation (Figure 2E–G, Figure 2—figure supplement 1E). Furthermore, nCa_V β was sufficient to confer greatly enhanced voltage-dependent inactivation to mCa_V (Figure 2E,G). From these results, we conclude that nCa_V β, the most enriched Ca_V subunit in nematocytes, confers nCa_V’s uniquely-sensitive voltage-dependent inactivation.

Because electrical stimulation has been implicated in nematocyte discharge and some nematocytes can produce action potentials (Anderson and Mckay, 1987; McKay and Anderson, 1988; Anderson and Bouchard, 2009), we used current-clamp to record the electrical responses of nematocytes to depolarizing stimuli. Under our conditions, nematocytes had a resting potential of −64.8 ± 8.9 mV and did not produce a voltage spike when injected with current from rest (Figure 3A). We further considered that the strong voltage-dependent inactivation of I_CaV could prevent excitability. Consistent with this idea, when nematocytes were first hyperpolarized to −90 mV and subsequently stimulated by current injection, we observed a singular long voltage spike (Figure 3B,C). In contrast, tentacle neurons produced multiple narrow spikes when injected with equivalent current amplitudes from a similar resting voltage, consistent with other neural systems (Figure 3A–C). Differences in spike width and frequency appear suited to mediate distinctive cellular functions: dynamic information processing in neurons and a single robust discharge event in nematocytes. Furthermore, these results indicate strong voltage-dependent inactivation prevents nematocyte activation from rest.

K⁺ channels contribute to resting membrane voltage (estimated reversal potential for nematocyte K⁺ is ~−100 mV) and often modulate repolarization following voltage spikes. Thus, we compared K⁺ currents in nematocytes and tentacle neurons to understand how spike width might be differentially regulated. Nematocytes exhibited transient outward K⁺ currents that quickly inactivated, while neurons had large sustained K⁺ currents, perhaps important for repolarization and repetitive spiking (Figure 3—figure supplement 1A–C). The transient component of the nematocyte K⁺ current was highly sensitive to voltage-dependent inactivation, similar to I_CaV (Figure 3—figure supplement 1D). This K⁺ current was abolished by the rapid intracellular Ca²⁺ chelator BAPTA or by removing external Ca²⁺, similar to the effect of the K⁺ channel blocker TEA⁺ (Figure 3—figure supplement 1E). Consistent with this observation, nematocyte-enriched cells expressed numerous Ca²⁺-activated K⁺ channels (Figure 3—figure supplement 1F). Furthermore, using intracellular Cs⁺ to block K⁺ currents resulted in prolonged voltage spikes and greatly increased resting membrane voltage, substantiating a role for K⁺ channels in modulating membrane voltage (Figure 3—figure supplement 1G). We propose that these distinct K⁺ channel properties could contribute to the singular wide spikes of nematocytes versus the numerous narrow spikes of neurons.

If nematocytes are basally inhibited due to the unique voltage-dependent inactivation of I_CaV, how do they respond to sensory signals to elicit discharge? Nematocyte discharge requires simultaneous detection of chemo- and mechanosensory cues (Pantin, 1942b; Watson and Hessinger, 1992), even though mechanical stimulation of the nematocyte’s cilium (cnidocil) within intact tentacles can by itself induce cellular depolarization (Brinkmann et al., 1996; Anderson and Bouchard, 2009). Indeed, we found the deflection of isolated nematocyte cnidocils elicited a mechanically gated inward current with rapid activation and inactivation kinetics. This current was abolished by gadolinium (Gd³⁺), which blocks mechanoreceptor and other cation channels, and was not observed in neurons (Figure 4A–C). Furthermore, nematocyte-enriched cells differentially expressed transcripts encoding NompC (no mechanoreceptor potential C, Figure 4—figure supplement 1A), a widely conserved mechanoreceptor previously found to localize to the cnidocil of nematocytes from Hydra (Schüler et al., 2015). Heterologous expression of Nematostella NompC (nNompC) resulted in a mechanically-gated current with similar properties to native nematocytes, including rapid kinetics and Gd³⁺ sensitivity (Figure 4A–C). Comparison with the Drosophila orthologue (dNompC) demonstrated nNompC had similar rapid kinetics, Gd³⁺ sensitivity, pressure-response relationships, and nonselective cation conductance, all consistent with the conservation of protein regions important for mechanosensitivity and ion selectivity (Jin et al., 2017; Figure 4A–C, Figure 4—figure supplement 1B–D). Thus, we conclude nematocytes are intrinsically mechanosensitive and suggest nNompC contributes to nematocyte mechanosensitivity. Importantly, this mechanically-evoked current is of sufficient amplitude to evoke a spike from very negative membrane voltages, but not from resting voltage at which I_CaV is inactivated.

Because nematocyte discharge is mediated by combined chemical and mechanical cues (Pantin, 1942b; Watson and Hessinger, 1992), we wondered if chemosensory signals could modulate nematocyte membrane voltage to allow for I_CaV activation and cellular responses. While prey-derived chemicals (<3 kDa extract from brine shrimp) evoked robust behavioral responses, similar treatments did not elicit electrical responses from isolated nematocytes (Figure 4D, Figure 4—figure supplement 1E). Considering in vivo cellular and discharge activity requires the presence of prey-derived chemicals with simultaneous mechanical stimulation, chemoreception may occur indirectly through functionally coupled cells (Price and Anderson, 2006). Previous studies suggest the presence of synaptic connections between nematocytes and other unknown cell types (Westfall et al., 1998; Oliver et al., 2008). To test this possibility, we screened isolated nematocytes for responses to well-conserved neurotransmitters and found only acetylcholine (ACh) elicited a significant response (Figure 4D,E). ACh-evoked outward currents were abolished by nicotinic acetylcholine receptor (nAChR) antagonists and recapitulated by nicotine (Figure 4F, Figure 4—figure supplement 1F). The K⁺ channel blocker TEA⁺ and the intracellular Ca²⁺ chelator BAPTA inhibited responses, suggesting ACh elicits K⁺ channel activity downstream of increased intracellular Ca²⁺. While the G-protein signaling inhibitor GDPβS did not affect outward currents, blockade of K⁺ currents with intracellular Cs⁺ revealed an ACh-elicited inward current that was enhanced by increased extracellular Ca²⁺ and blocked by the nAChR antagonist mecamylamine (Figure 4F, Figure 4—figure supplement 1G,H). These results suggest ACh evokes a Ca²⁺-permeable nAChR-like signaling pathway to engage Ca²⁺-activated K⁺ channels, consistent with the absence of muscarinic ACh receptors in Nematostella (Faltine-Gonzalez and Layden, 2019). In agreement with this observation, nematocyte-enriched cells expressed numerous nAChR-like transcripts which had well-conserved domains involved in Ca²⁺ permeability (Fucile, 2004; Figure 4—figure supplement 1I,J). Finally, we found robust acetylcholinesterase activity in tentacles, further suggesting a role for ACh signaling in nematocyte function (Figure 4G). These results demonstrate that nematocytes use cholinergic signaling to regulate K⁺ currents, similar to how efferent cholinergic innervation of vertebrate hair cells modulates nAChR-K⁺ channel signaling to inhibit auditory responses (Elgoyhen and Katz, 2012).

To identify the origin of cellular connections to nematocytes, we used serial electron microscopy reconstruction to visualize nematocytes and neighboring cells. We analyzed similar tentacle tissues from which we carried out physiological experiments and readily observed neurons and nematocytes in close proximity (Figure 5—figure supplement 1A,B). In resulting micrographs, nematocytes were clearly identified by their distinct nematocyst and cnidocil (Figure 5—figure supplement 1C). Interestingly, each nematocyte exhibited a long process, of presently unknown function, that extended into the ectoderm (Figure 5—figure supplement 1D). We also observed numerous spirocytes, indicated by the presence of a large intracellular thread-like structure (Figure 5A, Figure 5—figure supplement 1C). Putative sensory neurons were identified based on their synaptic contacts and extracellular projections (Figure 5A, Figure 5—figure supplement 1E,F). Importantly, dense core vesicles were localized to electron-dense regions at the junction between each nematocyte and one other cell type, either sensory neurons or spirocytes (Figure 5A, Figure 5—figure supplement 2A–E). Thus, nematocytes receive synaptic input from both neurons and spirocytes and likely serve as a site for integrating multiple signals (Figure 5—figure supplement 2F,G). This observation is consistent with the ability of cnidarians to simultaneously discharge multiple cnidocyte types to most efficiently capture prey (Pantin, 1942a).

How do distinct mechanosensory and chemosensory signals converge to elicit discharge? In agreement with our observation that nAChR activation increases K⁺ channel activity, ACh hyperpolarized nematocytes to negative voltages from which they were capable of producing robust voltage spikes (Figure 5B,C). A select number of cells with a more positive resting voltage still failed to produce spikes, and therefore additional regulation could exist through the modulation of resting membrane voltage (Figure 5—figure supplement 3A,B). These results suggest that the voltage-dependence for I_CaV prevents basal activation to depolarizing signals, such as mechanical stimulation, but activation of nAChR hyperpolarizes the cell to relieve I_CaV inactivation, thereby amplifying depolarizing signals to mediate cellular responses.

Consistent with a requirement for both mechano- and chemosensory input, we found the mechanoreceptor current blocker Gd³⁺ inhibited chemo-tactile stimulation of discharge. Additionally, the nAChR antagonist mecamylamine greatly reduced chemo-tactile-induced discharge (Figure 5D). Washout of both treatments recovered the ability of nematocytes to discharge (Figure 5—figure supplement 3C). Moreover, the requirement for prey-derived chemicals was completely recapitulated by ACh (Figure 5D). These results are consistent with a role for ACh signaling downstream of chemosensory stimulation. Thus, we propose that the unique I_CaV voltage-dependent inactivation provides a mechanism by which nematocytes filter extraneous depolarizing mechanical signals, but can integrate chemosensory-induced hyperpolarization together with a depolarizing stimulus to elicit robust signal amplification and discharge responses (Figure 6).

Starlet sea anemones (Nematostella vectensis) were provided by the Marine Biological Laboratory (Woods Hole, Massachusetts), Nv-Elav1::mOrange transgenic animals were a gift from F. Rentzsch. We used adult animals of both sexes, which were fed freshly-hatched brine shrimp (Artemia) twice a week and kept on a 12 hr light/dark cycle in 1/3 natural sea water (NSW). Nematocytes and neurons were isolated from tentacle tissue, which was harvested by anesthetizing animals in high-magnesium solution containing (mM): 140 NaCl, 3.3 Glucose, 3.3 KCl, 3.3 HEPES, 40 MgCl₂. Cells were isolated from tentacles immediately prior to electrophysiology experiments by treatment with 0.05% Trypsin at 32°C for 30 min and mechanical dissociation in divalent free recording solution (mM): 140 NaCl, 3.3 Glucose, 3.3 KCl, 3.3 HEPES, pH 7.6. Basitrichous isorhiza nematocytes were isolated from tentacles and identified by a thick-walled capsule containing a barbed thread, with a characteristic high refractive index, oblong shape and presence of a cnidocil. Spirocytes were identified by a thin-walled capsule containing a thin, unarmed thread, used for ensnaring prey. Neurons were identified by mOrange expression.

HEK293T cells (ATCC, Cat# CRL-3216, RRID:CVCL_0063, authenticated and validated as negative for mycoplasma by vendor) were grown in DMEM, 10% fetal calf serum, and 1% penicillin/streptomycin at 37°C, 5% CO₂. Cells were transfected using lipofectamine 2000 (Invitrogen/Life Technologies) according to the manufacturer’s protocol. 1 µg of Nematostella cacna1a, cacnb2.1, cacna2d1. M. musculus (mouse) cacna1a, R. norvegicus (rat) cacnb2a, or rat cacna2d1 was coexpressed with 0.5 µg GFP. Mechanosensitive proteins were assayed using HEK293T cells transfected with 1 µg of either Drosophila NompC or Nematostella NompC. To enhance channel expression, cells were transfected for 6–8 hr, plated on coverslips, and then incubated at 28°C for 2–6 days before experiments. Drosophila NompC-GFP was a gift from YN Jan. Rat cacna2d1 (RRID:Addgene_26575) and cacna1a (RRID:Addgene_26578) were gifts from D. Lipscombe and cacnb2a (RRID:Addgene_107424) was a gift from A. Dolphin.

RNA was prepared from tentacles of adult Nematostella using published methods (Stefanik et al., 2013). Briefly, 50 mg of tentacle tissue were homogenized and RNA was extracted using TRIzol. RNA was isolated and DNase treated (Zymo Research), then used for cDNA library synthesis (NEB E6560). Full-length sequence for a Nematostella calcium channel beta subunit was obtained with a RACE strategy using specific primers (GATTACGCCAAGCTTTATGCGTCCAATCGTACTTGTCGGC and GATTACGCCAAGCTTGCCGACAAGTACGATTGGACGCATA) on the amplified tentacle-tissue library. The final sequence was confirmed using primers corresponding to the end of the derived sequence (CAGAGCCAGGCCTGAGCGAG and GCCCCGTTAAAAGTCGAGAG) to amplify a full-length cDNA from tentacle mRNA, which was sequenced to confirm identity. nCa_v subunits: cacna1a, cacna2d1, cacnb2, and nNompC-GFP were synthesized by Genscript (Piscataway, NJ). Sequence alignments were carried out using Clustal Omega.

Tentacle tissue was ground to a fine powder in the presence of liquid nitrogen in lysis buffer (50 mM Tris-HCl pH 7.5, 250 mM KCl, 35 mM MgCl₂, 25 mM EGTA-KOH pH 8, 5 mM DTT, murine RNase inhibitor (NEB), 1% (w/v) NP-40, 5% (w/v) sucrose, 100 μg ml⁻¹ cycloheximide (Sigma), 500 μg ml⁻¹ heparin (Sigma)). Lysate was incubated on ice for 5 min, triturated five times with an 18 g needle, and insoluble material was removed by centrifugation at 16,000 g for 5 min at 4°C. Polyadenylated RNA was used to make sequencing libraries and sequenced on an Illumina HiSeq 4000 (Novogene). Quality filtering and adapter trimming was performed using Cutadapt (Martin, 2011), and a de novo transcriptome was assembled using Trinity (Grabherr et al., 2011). Annotation was performed using InterProScan (Jones et al., 2014) with Panther member database analysis, HMMER (Eddy, 2009) with the Pfam (El-Gebali et al., 2019) database, and DIAMOND (Buchfink et al., 2015) with the UniProt/TrEMBL database.

Reads from sorted cnidocytes (Bioproject PRJNA391807 Sunagar et al., 2018), enriched for nematocytes, were quality and adapter trimmed as described above, and transcript abundance (TPM) was quantified using Kallisto (Bray et al., 2016) and the tentacle transcriptome. For read mapping visualization, mapping was performed with Bowtie2 (Langmead and Salzberg, 2012), output files were converted to indexed bam files using Samtools (Li et al., 2009), and visualization was performed with the Integrated Genomics Viewer (Robinson et al., 2011).

Recordings were carried out at room temperature using a MultiClamp 700B amplifier (Axon Instruments) and digitized using a Digidata 1550B (Axon Instruments) interface and pClamp software (Axon Instruments). Whole-cell recording data were filtered at 1 kHz and sampled at 10 kHz. For single-channel recordings, data were filtered at 2 kHz and sampled at 20 kHz. Data were leak-subtracted online using a p/4 protocol, and membrane potentials were corrected for liquid junction potentials. For whole-cell nematocyte and neuron recordings, borosilicate glass pipettes were polished to 8–10 MΩ. The standard Nematostella medium was used as the extracellular solution and contained (in mM): 140 NaCl, 3.3 glucose, 3.3 KCl, 3.3 HEPES, 0.5 CaCl₂, 0.5 MgCl₂, pH 7.6. Two intracellular solutions were used for recording. For isolating inward currents (mM): 133.3 cesium methanesulfonate, 1.33 MgCl₂, 3.33 EGTA, 3.33 HEPES, 10 sucrose, 10 CsEGTA, pH 7.6. For outward currents (mM) 166.67 potassium gluconate, 3.33 HEPES, 10 sucrose, 1.33 MgCl₂, 10 KEGTA, pH 7.6. In some experiments, BAPTA was substituted for EGTA. For whole-cell recordings in HEK293 cells, pipettes were 3–4 MΩ. The standard extracellular solution contained (in mM): 140 NaCl, 5 KCl, 10 HEPES, 2 CaCl₂, 1 MgCl₂, pH 7.4. The intracellular solution contained (mM): 140 cesium methanesulfonate, 1 MgCl₂, 3.33 EGTA, 3.33 HEPES, 10 sucrose, pH 7.2. In ion substitution experiments, equimolar Ba²⁺ was substituted for Ca²⁺. Single-channel recording extracellular solution contained (mM): 140 NaCl, 10 HEPES, 1 NaEGTA, pH 7.4. The intracellular solution used (mM): 140 CsCl, 10 HEPES, 1 CsEGTA, pH 7.4.

The following pharmacological agents were used: N-Acetylneuraminic acid (NANA, 100 µM, Sigma), glycine (100 µM, Sigma), acetylcholine (1 mM), mecamylamine (100 µM, 500 µM for behavioral experiments, Tocris), GdCl₃ (100 µM, Sigma), glutamate (1 mM), GABA (1 mM), nicotine (100 µM, Tocris), tubocurarine (10 µM, Tocris), TEA⁺ (10 mM, Sigma), BAPTA (10 mM, Tocris), GDPβS (1 mM, Sigma), and Cd²⁺ (500 µM, 250 µM for behavioral experiments). All were dissolved in water. Prey extract was isolated from freshly-hatched Artemia. Artemia were flash-frozen, ground with mortar and pestle, filtered with 0.22 µM pores or 3 kDa ultracentrifugal filters (Amicon UFC500324). Pharmacological effects were quantified as differences in normalized peak current from the same cell following bath application of the drug (I_treatment/I_control). Whole-cell recordings were used to assess mechanical sensation together with a piezoelectric-driven (Physik Instrumente) fire-polished glass pipette (tip diameter 1 μm). Mechanical steps in 0.5 μm increment were applied every 5 s while cells were voltage-clamped at −90 mV. Single mechanosensitive channels were studied using excised outside-out patches exposed to pressure applied via a High-Speed Pressure Clamp system (HSPC, ALA-scientific). Pressure-response relationships were established using pressure steps in 10 mmHg increments. Voltage-dependence of currents was measured from −100 mV to 100 mV in 20 mV increments while applying repetitive 60 mmHg pressure pulses.

Unless stated otherwise, voltage-gated currents were measured in response to a 200 ms voltage pulse in 10 mV increments from a −110 mV holding potential. G-V relationships were derived from I-V curves by calculating G: G = I_CaV/(V_m-E_rev) and fit with a Boltzman equation. Voltage-dependent inactivation was measured during −10 mV (Ca²⁺ currents in native cells), 0 mV (Ca²⁺ currents in heterologously expressed channels), 60 mV (K⁺ currents in native cells) voltage pulses following a series of 1 s pre-pulses ranging from −110 mV to 60 mV. Voltage-dependent inactivation was quantified as I/I_max, with I_max occurring at the voltage pulse following a −110 mV prepulse. In some instances, inactivation curves could not be fit with a Boltzman equation and were instead fitted with an exponential. The time course of voltage-dependent inactivation was measured by using a holding voltage of −110 mV or −70 mV and applying a 0 mV test pulse every 5 s. Recovery from inactivation was quantified by normalizing inactivated and recovered currents to those elicited from the same cell in which a 0 mV voltage pulse was applied from −110 mV. Test pulses from a holding voltage of −40 mV were used to assess inactivation, followed by pulses from −110 mV where currents quickly recovered to maximal amplitude. Repetitive stimulation using 20 ms pulses to −10 mV or 0 mV from a holding voltage of −90 mV was also used to measure inactivation in response to repetitive stimulation. Current inactivation kinetics were quantified by the portion of current remaining at the end of a 200 ms pulse (R200) or fit with a single exponential. Activation was quantified as the time from current activation until peak amplitude. 200 ms voltage ramps from −120 to 100 mV were used to measure ACh-elicited currents. Stimulus-evoked currents were normalized to basal currents measured at the same voltage of 80 mV.

Single channel currents were measured from the middle of the noise band between closed and open states or derived from all-points amplitude histograms fit with Gaussian relationships at closed and open peaks for each excised patch record. Conductance was calculated from the linear slope of I–V relationships. N(P_O) was calculated during pressure steps while voltage was held at −80 mV. In current clamp recordings, effects of ACh or intracellular ions on resting membrane potential was measured without current injection (I = 0). 1 s depolarizing current steps of various amplitudes were injected to measure spikes which were quantified by frequency (spikes/second) or width (duration of spike). To test whether resting membrane potential affects the ability to generate spikes, hyperpolarizing current was injected to bring cells to negative voltages (<-90mV) or ACh was locally perfused before depolarizing current injection.

The change in Ca²⁺ concentration from a nematocyte voltage spike was estimated based on the integral of Ca²⁺-selective nematocyte currents elicited by a 0 mV step, the same amplitude and slightly shorter duration than a voltage spike. Nematocyte volume was estimated from serial electron microscopy reconstruction with a non-nematocyst volume of approximately 5% of the total volume of the cell. We did not consider the volume occupied by other organelles, making for a conservative estimate. Furthermore, calculations were made with extracellular recording solution containing 0.5 mM Ca²⁺, which is approximately six-fold less than physiological concentrations. Thus, the large increase we calculated likely underestimates the total Ca²⁺ influx.

Discharge of nematocysts was assessed based on well-established assays (Watson and Hessinger, 1994a; Gitter et al., 1994). Adult Nematostella were placed in petri dishes containing a modified Nematostella medium, containing 16.6 mM MgCl₂. Animals were given appropriate time to acclimate before presented with stimuli. For assaying discharge, 5 mm round coverslips were coated with a solution of 25% gelatin (w/v) dissolved in medium, and allowed to cure overnight prior to use. Coverslips were presented to the animal’s tentacles for 5 s and then immediately imaged at 20X magnification using a transmitted light source. To assay behavioral response to prey-derived chemicals, freshly hatched brine shrimp were flash frozen and pulverized, then filtered through a 0.22 µm filter. Coverslips were dipped in the prey extract and immediately presented to the animal. All pharmacological agents were bath applied, except for acetylcholine (1 mM), which was delivered as a bolus immediately prior to coverslip presentation. Acetylcholine exposure did not produce movement or contraction of tentacles. Experiments carried out in the absence of extracellular Ca²⁺ were nominally Ca²⁺ free and did not use extracellular chelators. The highest density of discharged nematocytes on the coverslip was imaged at 20X. Nematocyte discharge involves everting the barbed thread, causing them to embed in the gelatin-coated coverslips. Therefore, if nematocytes do not discharge, they are not captured by the gelatin-coated coverslip or visualized for quantification. Images were blindly analyzed using a custom Matlab routine (available in supplemental material) in which images were thresholded and the fraction of pixels corresponding to nematocytes was compared across experiments.

Tentacles from an individual Nematostella vectensis were placed between two sapphire coverslips separated by a 100 μm spacer ring (Leica) and frozen in a high-pressure freezer (EM ICE, Leica). This was followed by freeze-substitution (EM AFS2, Leica) in dry acetone containing 1% ddH2O, 1% OsO4 and 1% glutaraldehyde at −90°C for 48 hr. The temperature was then increased at 5°C/h up to 20°C and samples were washed at room temperature in pure acetone 3 × 10 min RT and propylene oxide 1 × 10 min. Samples were infiltrated with 1:1 Epon:propylene oxide overnight at 4°C. The samples were subsequently embedded in TAAB Epon (Marivac Canada Inc) and polymerized at 60°C for 48 hr. Ultrathin sections (about 50 nm) were cut on an ultramicrotome (Leica EM UC6) and collected with an automated tape collector (ATUM Kasthuri et al., 2015). The sections were then post-stained with uranyl acetate and lead citrate prior to imaging with a scanning electron microscope (Zeiss SIGMA) using a back-scattered electron detector and a pixel size of 4 nm.

Once all the sections were scanned, images were aligned into a stack using the algorithm ‘Linear Stack Alignment with SIFT’ available in Fiji (Schindelin et al., 2012). After alignment, images were imported into VAST (Berger et al., 2018) so that every cell could be manually traced. By examining sections and following cellular processes contacts between cells of interest (e.g. neurons and nematocytes) were identified and assessed for the presence of dense-core vesicles in the vicinity (~500 nm). Such instances were labeled as putative synapses. Cells were then rendered in three dimensions using 3D Studio Max 2019 (Autodesk, San Rafael, CA).

Nematocytes were readily identified because resin does not infiltrate the nematocyst capsule, making for an ‘empty’ appearance (large white area). Spirocytes were also readily identified based on their capsule containing a long, coiled filament. Sensory neurons were identified according to the higher number of dense core vesicles, higher number of synapses and the presence of sensory processes extending into the external environment.

Data were analyzed with Clampfit (Axon Instruments) or Prism (Graphpad) and are represented as mean ± s.e.m. n represents independent experiments for the number of cells/patches or behavioral trials. Data were considered significant if p<0.05 using paired or unpaired two-tailed Student’s t-tests or one- or two-way ANOVAs. All significance tests were justified considering the experimental design and we assumed normal distribution and variance, as is common for similar experiments. Sample sizes were chosen based on the number of independent experiments required for statistical significance and technical feasibility.