We recently reported the synthesis of the β‐scorpion toxin Cn2, a highly potent and selective activator of Na V 1.6 channels (Schiavon et al . 2006 ; Israel et al . 2018 ). Equipped with Cn2 we sought to explore the effects of Na V 1.6 channel activation in peripheral sensory neurons using a multidisciplinary approach of patch‐clamp recordings, recordings in isolated skin–nerve and gut–nerve preparations, calcium imaging and animal behaviour studies. Our data confirm the selectivity of Cn2 in native neurons and support an important role for Na V 1.6 channels in sensory neurons that is crucial for painful mechanosensory transduction in the skin and gut.

There are several hurdles for studying native Na V 1.6 currents in peripheral sensory neurons in adult animals. The macroscopic TTX‐S currents produced by Na V 1.1, Na V 1.6 and Na V 1.7 cannot be readily distinguished especially as the relative of each of these channels to the total TTX‐S current is not well understood. The use of Na V 1.6 global knock‐out mice for behavioural studies is limited to animals that are less than 2 weeks old because Na V 1.6 −/− ( SCN8A medtg/medtg ) mice begin to show stunted growth by P12 and are juvenile lethal (Burgess et al . 1995 ). This phenotype also precludes recordings from sensory afferents using the murine skin–saphenous nerve preparation. Furthermore, isolating Na V 1.6 pharmacologically has proved difficult. The TTX analogue 4,9‐anhydro‐TTX has been reported to display high selectivity for Na V 1.6 over other TTX‐S isoforms (Rosker et al . 2007 ), albeit more recent studies failed to replicate high‐potency inhibition of Na V 1.6 (Tsukamoto et al . 2017 ). Thus, novel approaches are needed to study Na V 1.6 channels in isolation in native neurons.

Evidence from murine models confirmed a role for Na V 1.6 channels in the development and maintenance of oxaliplatin‐induced cold allodynia and neuropathic pain after spinal nerve ligation (Sittl et al . 2012 ; Deuis et al . 2013 ; Xie et al . 2015 ), and inflammation of dorsal root ganglion (DRG) tissue (Xie et al . 2013 ). Conditional knock‐out of Na V 1.6 channels in adult small‐ and large‐diameter neurons reduces neuropathic pain associated with spared nerve injury (Chen et al . 2018 ). Additionally, a gain‐of‐function mutation in Na V 1.6 was recently identified in a patient with trigeminal neuralgia, causing hyper‐excitability of trigeminal neurons (Tanaka et al . 2016 ). Although this evidence suggests that Na V 1.6 plays a crucial role in peripheral neuronal activity, the precise contribution of Na V 1.6 channels to excitability of sensory neurons remains unclear, and a systematic assessment of Na V 1.6 channel function in these neurons is required for a comprehensive understanding of the contribution of this channel to pain disorders.

Voltage‐gated sodium channels (Na V ) are responsible for the upstroke of the action potentials in electrically excitable cells (Hille, 2001 ). In total, there are nine Na V subtypes (Na V 1.1–Na V 1.9) with different functional properties, and a subset is expressed at substantial levels in adult peripheral sensory neurons. Anatomically, peripheral sensory neurons are pseudo‐unipolar with a central terminus projecting to the dorsal horn of the spinal cord and a projection terminating in peripheral tissue, including the skin and viscera. Peripheral Na V subtypes include the tetrodotoxin‐sensitive (TTX‐S) Na V 1.1, Na V 1.6 and Na V 1.7 channels and the TTX‐resistant (TTX‐R) Na V 1.8 and Na V 1.9 channels. Na V 1.7–Na V 1.9 channels are preferentially expressed in peripheral neurons and have been associated with a range of painful and painless channelopathies (Dib‐Hajj et al . 2013 , 2015 ; Erickson et al . 2018 ). Na V 1.6 channels are expressed in both the central and peripheral nervous system where they are the major Na V isoform at the nodes of Ranvier and are also present in unmyelinated fibres and at the nerve terminals of a subset of sensory neurons (Caldwell et al . 2000 ; Persson et al . 2010 ; Carrasco et al . 2017 ; Chen et al . 2018 ). Indeed, global loss of Na V 1.6 reduced the compound action potential of both A‐ and C‐fibres (Black et al . 2002 ). The fast kinetics of Na V 1.6 channels, including rapid repriming (Herzog et al . 2003a ), and the production of relatively large persistent and resurgent current (Cummins et al . 2005 ; Rush et al . 2005 ) allow for high‐frequency firing in sensory neurons that express this isoform (Herzog et al . 2003a ).

The data and statistical analysis in this study comply with the recommendations on experimental design and analysis in pharmacology (Curtis et al . 2018 ). Unless otherwise noted, statistical significance was determined using paired Student's t test. Results are presented as mean ± SEM, and P < 0.05 was considered to show a statistically significant difference between Control and Cn2‐treated conditions. Statistics were carried out using Prism (version 7).

The effect of Cn2 on TRPA1 and TRPM8 channels was assessed using a high‐throughput fluorescence Ca 2+ assay as previously described (Vetter et al . 2012 ). In brief, HEK293 cells stably expressing TRPA1 and TRPM8 channels were plated on clear‐bottom 384‐well plates at a density of 15,000 cells per well and loaded with Calcium 4 No‐Wash dye (Molecular Devices, Sunnyvale, CA, USA) diluted in physiological salt solution (buffer; composition n mM: 140 NaCl, 11.5 glucose, 5.9 KCl, 1.4 MgCl 2 , 1.2 NaH 2 PO 4 , 5 NaHCO 3 , 1.8 CaCl 2 and 10 Hepes) for 30 min at 37°C. Fluorescence responses to addition of Cn2 (10 nM, 100 nM and 1 µM) were measured every second for 300 s using a FLIPR TetraPlus fluorescence plate reader (excitation 470–495 nm; emission 515–575 nm; Molecular Devices) followed by stimulation with menthol (1 mM) in TRPM8‐expressing cells or allyl isothiocyanate (1 mM) in TRPA1‐expressing cells. Responses were analysed using ScreenWorks 3.2.0.14 (Molecular Devices) and plotted in Prism (version 7).

All animal behavioural experiments utilized male C57BL/6 mice, 6–8 weeks, approximately 20–25 g. After experimentation all animals were sacrificed by CO 2 inhalation. To assess the effect of Na V 1.6 channel activation in vivo , a previously described model of Cn2‐evoked nociception was utilized (Deuis et al . 2013 ). Under brief isoflurane (3%) anaesthesia, a single dose of Cn2 (1–10 nM) was diluted in sterile PBS containing 0.1% BSA and administered via shallow subcutaneous injection to the hindpaw (intraplantar, 40 µL, 10 nM). Animals were placed in clear Perspex boxes to recover and recorded for 1 h post‐injection to assess spontaneous pain. Following cessation of spontaneous pain behaviours (this includes flinches, lifts, licks and shakes of the hindpaw), mechanical allodynia was tested using the MouseMet ™ electronic von Frey (TopCat Metrology, Little Downham, UK). Thermal threshold was assessed with the MouseMet ™ thermal probe (TopCat Metrology).

To assess the effects of Cn2 on colonic nociceptor mechanosensitivity, we performed single afferent recordings from splanchnic nerves as described (Osteen et al . 2016 ; Castro et al . 2017 , 2018 ; Inserra et al . 2017 ). Briefly, the colon and rectum (5–6 cm) was removed from C57BL/6 mice. Subsequently, the tissue was opened and pinned flat, mucosal side up, in an organ bath. Tissue was superfused with a modified Krebs solution (mM: 117.9 NaCl, 4.7 KCl, 25 NaHCO 3 , 1.3 NaH 2 PO 4 , 1.2 MgSO 4 .7H 2 O, 2.5 CaCl 2 and 11.1 d‐glucose, 95% O 2 –5% CO 2 , 34°C), containing the L‐type calcium channel antagonist nifedipine (1 µM) to suppress smooth muscle activity, and the cyclooxygenase inhibitor indomethacin (3 µM) to suppress inhibitory actions of prostaglandins. The splanchnic nerve was extended into a paraffin‐filled recording compartment, in which finely dissected strands were laid onto a platinum electrode for single‐unit extracellular recordings of action potentials generated by mechanical stimulation of receptive fields in the colon. Receptive fields were identified by mechanical stimulation of the mucosal surface with a brush of sufficient stiffness to activate all types of mechanosensitive afferents. Once identified, receptive fields were tested with three distinct mechanical stimuli to enable classification: static probing with calibrated von Frey filaments (2 g force; 3 times for 3 s), mucosal stroking with von Frey filaments (10 mg force; 10 times), or circular stretch (5 g; 1 min) (Castro et al . 2013 , 2017 ; Bellono et al . 2017 ). In this study, we focused on high‐threshold colonic nociceptive afferents, also termed serosal or vascular afferents. These fibres responded to noxious distension (40 mmHg), circular stretch (≥7 g) or 2 g filament probing, but not to fine mucosal stroking (10 mg filament), and they express an array of channels and receptors involved in pain, become mechanically hypersensitive in models of chronic visceral pain, and have a nociceptor phenotype (Erickson et al . 2018 ; Sadeghi et al . 2018 ). In the current study, they are therefore referred to as ‘colonic nociceptors’. Once baseline colonic nociceptor responses to mechanical stimuli (2 g von Frey hair filament) had been established, mechanosensitivity was re‐tested after 5 min application of the β‐scorpion toxin Cn2 (1 µM). Cn2 was applied to the mucosal surface of the colon via a small metal ring placed over the receptive field of interest. This route of administration has been shown to activate colonic afferents (Osteen et al . 2016 ; Inserra et al . 2017 ; Jimenez‐Vargas et al . 2018 ). Action potentials were analysed using the Spike2 wavemark function (Cambridge Electronic Design Limited, Cambridge, UK) and discriminated as single units on the basis of distinguishable waveform, amplitude and duration.

To assess the pharmacology of toxins on sensory A‐ and C‐fibre afferents, we utilized the skin–saphenous nerve preparation as previously described (Reeh, 1986 ). Juvenile male Wistar rats (200–250 g) were sacrificed by CO 2 inhalation and the hairy skin of the hindpaws and legs was dissected with the saphenous nerve attached. The first skin of each pair was placed in a recording chamber, secured hairy side down with petroleum jelly and continuously perfused with carbogenated synthetic interstitial fluid (SIF) composed of (mM): 107.8 NaCl, 3.5 KCl, 0.69 MgSO 4 .2H 2 O, 26.2 NaHCO 3 ,1.67 NaH 2 PO 4 .2H 2 O, 9.64 gluconic acid, 5.55 glucose, 7.6 sucrose and 1.53 CaCl 2 .2H 2 O, pH 7.3; the second skin of the animal was kept at 4°C until ready for experimentation (up to 6 h post‐dissection). The saphenous nerve bundle of the first skin was then placed on a mirror in a separate recording chamber filled with paraffin oil, de‐sheathed and teased to smaller filaments which were placed on a platinum recording electrode. The corium of the skin was then manually probed with a blunt glass rod until single, mechanically sensitive receptive fields could be identified and subsequently classified by conduction velocity (C < 1 m/s; A > 1.6–12 m/s) after stimulation by bipolar Teflon‐coated steel microelectrode (impedance 1\MΩ). Mechanical thresholds were determined by a gravity‐driven von Frey probe. The receptive field was then isolated using a small plastic ring and baseline response to heat (>50°C) and cold (<10°C) stimulus was assessed by perfusion of temperature‐controlled SIF. Toxin was then perfused for 5 min. Subsequently, C‐fibre receptive fields were subjected to temperature ramps and the pre‐determined threshold mechanical stimulus was applied. For A‐fibres, the threshold mechanical stimulus was applied briefly (5 s stimulus) and resulting action potential generation recorded using DAPSYS (Brian Turnquist; www.dapsys.net ) software.

Current‐clamp recordings were obtained using a HEKA EPC‐10 double amplifier (acquired at 50 kHz and filtered at 2.9 kHz with a low‐pass Bessel filter). ECS contained (mM); 140 NaCl, 3 KCl, 2 MgCl 2 , 2 CaCl 2 , 10 Hepes and 10 dextrose (pH and osmolarity adjusted as above). Intracellular solution contained (mM): 140 KCl, 0.5 EGTA, 5 Hepes and 10 dextrose (pH adjusted to 7.3 with KOH, osmolarity adjusted to 310 with sucrose). Patch pipettes (1–2 MΩ resistance) were pulled, using a Sutter Instrument Co. P‐97, from borosilicate glass (1.65/1.1, OD/ID; World Precision Instruments) and fire polished (Microforge MF830, Narishige). Whole‐cell configuration of large‐diameter DRG neurons (>35 µm) were obtained in voltage‐clamp prior to proceeding to current‐clamp mode. Cells were continuously perfused with ECS (in 0.1% BSA) in control recordings. Cells with a stable resting membrane potential (RMP), recorded for 30 s and found to have an RMP ≤−55 mV were recorded. Input resistance was determined by hyperpolarizing current steps at 10 pA increments between −10 and −60 pA and fitted to linear regression. Current threshold was determined to be the first action potential elicited after serial depolarizing current injections in 10 pA increments. Repetitive firing was assessed by increasing current injection in 100 pA increments up to two times the current threshold. Action potential amplitude was measured from the action potential peak to RMP. All current‐clamp parameters were tested post perfusion (2–3 min) with Cn2 in ECS (0.1% BSA). Data were analysed offline with Fitmaster (HEKA Electronik), Microsoft Excel and plotted using Prism software (version 7, GraphPad Software Inc., La Jolla, CA, USA).

Protocols included a prepulse step to 0 mV to allow toxin binding as per the voltage‐sensor trapping model, and repolarization to −100 mV for 100 ms to allow recovery from fast inactivation (Cestele et al . 1998 ; Schiavon et al . 2006 ). This was followed by measurement of current–voltage ( I–V ) relationships obtained from sweeps between −80 mV and +20 mV for 100 ms at 5 s intervals. Following control recording, ECS containing toxins (in 0.1% BSA) was perfused for 2–3 min through a pressure driven perfusion system with a perfusion pencil (AutoMate Scientific, Berkeley, CA, USA) to allow sufficient fluid exchange and toxin binding, which was followed by recording with continuous perfusion.

Whole‐cell electrophysiology recordings were obtained from mouse DRG neurons using an EPC‐10 double amplifier and Patchmaster (HEKA Elektronik, Holliston, MA, USA) software. Voltage‐clamp recordings were undertaken at room temperature 20 ± 1°C using an extracellular solution (ECS) containing (mM): 30 NaCl, 110 choline‐Cl, 3 KCl, 1 MgCl 2 , 1 CaCl 2 , 10 Hepes, 5 CsCl, 20 tetraethylammonium‐Cl, 0.1 CdCl 2 and 1 4‐aminopyridine, pH 7.3 adjusted with NaOH, osmolarity 315−320 mOsm adjusted with dextrose. Intracellular recording solution contained (mM): 130 CsMeSO 4 , 20 NaCl, 0.2 EGTA, 10 Hepes, 4 Mg‐ATP, 0.3 Na‐GTP and 10 dextrose, pH 7.3 adjusted with CsOH, osmolarity 300–305 mOsm adjusted with dextrose. Patch pipettes (0.7–1.3 MΩ resistance) were pulled from borosilicate glass (1.65/1.1, OD/ID, World Precision Instruments, Sarasota, FL, USA) using a Sutter Instrument Co. (Novato, CA, USA) P‐97 puller and fire polished (Microforge MF830, Narishige, Amityville, NY, USA). Whole‐cell configuration was achieved and DRG neurons were held at −80 mV. Voltage error was kept below ±10 mV for medium and large‐diameter DRG neurons, and ±5 mV for small‐diameter DRG neurons using 70–90% series resistance compensation. Recordings were acquired at 50 kHz and filtered with a low‐pass Bessel filter at 2.9 kHz, and linear leak was corrected by P /6 subtraction.

DRG neurons were loaded with Fluo‐4 AM calcium indicator according to the manufacturer's instructions (Thermo Fisher Scientific), and incubated for 1 h at 37°C (Robinson et al . 2018 ). After loading (1 h), the dye‐containing solution was replaced with assay solution (1× Hanks’ balanced salt solution, 20 mM Hepes). Fluorescence corresponding to [Ca 2+ ] i of typically 100–150 DRG cells per experiment was monitored in parallel using an Nikon Ti‐E Deconvolution inverted microscope, equipped with a Lumencor Spectra LED light source. Images were acquired through a 20× objective at 1 frame/s (excitation 485 nm, emission 521 nm). For each experiment, baseline fluorescence was monitored for 30 s. At 30, 60, 90 and 150 s, assay solution was replaced with assay solution (negative control), Cn2 (500 nM in assay solution), assay solution (wash) and KCl (30 mM in assay solution), respectively. Cells responding to KCl and/or Cn2 were considered neuronal and grouped according to size: large (>600 µm 2 ), medium (300–600 µm 2 ) and small diameter (<300 µm 2 ).

DRG neuron isolation for calcium imaging was performed as previously reported (Teichert et al . 2012 ; Robinson et al . 2018 ). DRGs (all spinal levels) from 4‐ to 8‐week‐old male C57BL/6 mice were dissociated with collagenase, (1 mg/mL for 90 min at 37°C) and plated in DMEM (ThermoFisher Scientific) containing 10% fetal bovine serum (FBS) (Assay Matrix, Melbourne, VIC, Australia) and penicillin/streptomycin (ThermoFisher Scientific) on a 96‐well poly‐d‐lysine‐coated culture plate (Corning, Tewksbury, MA, USA) and maintained for 20–24 h at 37°C, 5% CO 2 before assay.

DRG neurons from thoracic and lumbar regions were isolated from Nav1.6 −/− ( Scn8a medtg/medtg ) mice (P11–P17, both male and female) or C57BL/6 mice (4–8 weeks old, both male and female) as previously reported (Dib‐Hajj et al . 2009 ). Briefly, DRGs were harvested from homozygous mice, incubated at 37°C for 20 min in complete saline solution (CSS) (in mM: 137 NaCl, 5.3 KCl, 1 MgCl 2 , 25 sorbitol, 3 CaCl 2 and 10 Hepes, adjusted to pH 7.2 with NaOH) containing 0.5 U/mL Liberase TM (Sigma Aldrich, St. Louis, MO, USA) and 0.6 mM EDTA, followed by a 15 min incubation at 37°C in CSS containing 0.5 U/mL Liberase TL (Sigma‐Aldrich), 0.6 mM EDTA, and 30 U/mL papain (Worthington Biochemical Corp., Lakewood, NJ, USA). DRGs were then centrifuged and triturated in 0.5 mL of DRG medium: Dulbecco's modified Eagle's medium–F12 (1:1) with 100 U/mL penicillin, 0.1 mg/mL streptomycin (ThermoFisher Scientific, Waltham, MA, USA), and 10% fetal bovine serum (GE Healthcare Bio‐Sciences, Pittsburgh, PA, USA), containing 1.5 mg/mL BSA (low endotoxin; Sigma‐Aldrich) and 1.5 mg/mL trypsin inhibitor (Sigma‐Aldrich). After trituration, the cells were diluted with DRG medium containing 1.5 mg/mL BSA and 1.5 mg/mL trypsin inhibitor, seeded onto poly‐d‐lysine/laminin‐coated coverslips (BD Biosciences, San Jose, CA, USA), and incubated at 37°C in a 95% air–5% CO 2 (v/v) incubator for 45 min for neurons to attach to the coverslips. After 45 min, DRG medium was added into each well to a final volume of 1 mL and the DRG neurons were maintained at 37°C in a 5% CO 2 (v/v) incubator for 18–30 h before current‐clamp or voltage‐clamp recording. For current‐clamp recordings, culture medium was supplemented with 50 ng/mL mouse nerve growth factor (mNGF) (Alomone Labs, Jerusalem, Israel) and 50 ng/mL recombinant human glial cell line‐derived neurotrophic factor (hGDNF) (PeproTech, Rocky Hill, NJ, USA).

All animal experiments were approved by the University of Queensland, the South Australian Health and Medical Research Institute (SAHMRI) or the Flinders University animal ethics committees and were conducted in accordance with the NHMRC code for use of animals (2013 edition) and the International Association for the Study of Pain Guidelines for the Use of Animals in Research. All experiments performed conformed to the relevant regulatory standards and the ARRIVE guidelines (McGrath & Lilley, 2015 ). Animal studies followed a protocol approved by the Department of Veterans Affairs West Haven Medical Centre (VAMC) Animal Use Committee. C57BL/6 mice and Wistar rats were housed in the University of Queensland School of Pharmacy (PACE), South Australian Health and Medical Research Institute (SAHMRI), and the Centre for Neuroscience and Regeneration Research, VAMC, West Haven, CT, USA. Mice were housed in individually ventilated cages (≤5 animals per cage) with corn cob bedding or aspen wood chip bedding. Cage racks were in temperature‐controlled rooms (22°C) and exposed to a 12 h light–12 h dark cycle. Animals were given access to standard rodent chow and water ad libitum and supplied with a red polycarbonate Mouse House (Techniplast, Italy) and shredded paper for nesting and enrichment. All animals were acclimatized (for 1 h) in the behavioural room prior to behavioural experiments.

After injection of Cn2 (10 nM) into the dorsal hindpaw (intraplantar, in 0.1% BSA), Cn2 causes spontaneous pain quantified by the number of lifts, licks and shakes of the hindpaw. A , time course of non‐stimulus evoked nocifencive behaviours post‐Cn2 (10 nM) injection. B , total number of flinches, flicks, licks and shakes of hindpaw at peak, 5 min after administration (Control: 1 ± 1; Cn2: 72.7 ± 10.1). C , after cessation of all spontaneous pain behaviours (>30 min post‐administration), Cn2 causes a reduction in paw withdrawal threshold to von Frey filament (Control: 2.3 ± 0.3, n = 13; Cn2: 1.0 ± 0.2, n = 8; P < 0.05, unpaired t test). C , intraplantar injection of Cn2 (10 nM) does not affect noxious temperature sensing (paw withdrawal temperature: Control: 50.6 ± 0.9°C; Cn2: 50.5 ± 1.3°C; P > 0.05, unpaired t test).

Consistent with the bursting phenotype observed in a subset of A‐fibres, as well as previous reports (Deuis et al . 2013 ; Israel et al . 2018 ), Cn2 induced spontaneous pain when administered by the intra‐plantar route (Fig. 9 A and B ), quantified by the number of lifts, licks, shakes and flinches of the hindpaw. Spontaneous nocifencive behaviours persist for 30 min (Fig. 9 A ). Following cessation of spontaneous pain behaviours (>30 min post‐injection), Cn2 (10 nM) caused a significant decrease in the mechanical paw withdrawal threshold compared to vehicle control (Fig. 8 C ). Consistent with the limited effects observed in C‐fibres in ex vivo recordings, Cn2 did not affect behavioural responses to cold stimuli (Deuis et al . 2013 ). In addition, the heat withdrawal threshold remained unchanged between control and Cn2‐treated animals (Fig. 9 D ).

A , representative trace of splanchnic nerve recording showing that Cn2 (1 µM, 5 min) failed to evoke colonic nociceptor firing. B , representative traces of colonic nociceptor mechanosensitivity before and after Cn2 (1 µM, 5 min) using 2 g von Frey filaments (vfh) applied to the receptive field of the colon. C , increase in number of action potentials per second in response to mechanical stimulus (Control: 10.8 ± 0.8; Cn2: 15.7 ± 1.3; n = 16, P < 0.01, unpaired sample t test). D , increase in the peak firing frequency, in response to mechanical stimulus (Control: 20.9 ± 2.7 Hz; Cn2: 34.1 ± 3.0 Hz; n = 16, P < 0.05, unpaired sample t test).

We also investigated the functional role of Na V 1.6 channels at peripheral terminals in the viscera using the colon–splanchnic nerve preparation. All of these colonic afferents have conduction velocities within the C‐fibre range (<2 m/s) (Jones et al . 2005 ). Application of Cn2 to the mucosal surface of the colon did not evoke spontaneous firing in any of the colonic nociceptors (C‐fibres) tested (Fig. 8 A ). However, 5 min incubation with Cn2 (1 µM) subsequently resulted in a significant increase in both the number of action potentials per second and the peak firing in response to mechanical stimuli (Fig. 8 B and C ).

A , representative trace of C‐fibre mechano‐heat (CMH) fibre. B , pooled peak frequency of firing from heat sensitive C fibres showing non‐significant increase in firing in response to 10 nM Cn2 ( n = 6, P > 0.05, paired t test). C , representative trace C‐fibre mechano‐cold (CMC) fibre types. D , cold sensitive C‐fibres display a small decrease in number of action potentials after Cn2 (10 nM) application (Control: 51.9 ± 4.6; Cn2: 31.5 ± 5.7; n = 13, P < 0.05, paired t test). E , representative response to grams of stimulus (g) after Cn2 (10 nM) showing no increase in peak frequency or number of action potentials. F , pooled peak frequency of firing at threshold of mechano‐sensitive C fibres before and after Cn2 application (Control: 32.3 ± 11.0/s; Cn2: 33.6 ± 11.0/s).

Subtle changes to firing responses were observed in C‐fibres (in the skin) in the presence of Cn2 (Fig. 6 ). The conduction velocity of skin‐afferent C‐fibres ranged from 0.1 to 0.9 m/s, in line with previous reports (CV: 0.47 ± 0.08 m/s) (Zimmermann et al. 2009 ). Cn2 did not significantly enhance action potential firing of heat‐sensitive C‐fibres (Fig. 6 A and B ). By contrast, cold‐mechanosensitive C‐fibres displayed a significant decrease in the number of action potentials in response to cold stimuli (Fig. 6 C ). To exclude Cn2‐mediated effects on cold‐sensitive transient receptor potential (TRP) channels, we assessed Ca 2+ responses to Cn2, as well as effects of Cn2 on allyl isothiocyanate (AITC) and menthol responses, in TRPA1‐ and TRPM8‐expressing HEK293 cells. Consistent with selective activity at Na V 1.6, Cn2 neither activated TRPA1 or TRPM8, nor modulated AITC‐ or menthol‐induced TRPA1 and TRPM8 responses (Fig. 7 A–D ). In addition, C‐fibres did not spontaneously fire after Cn2 application and did not have increased responses to mechanical stimulus (Fig. 6 D and E ).

A and B , representative traces of A‐fibre mechanosensitivity before and after Cn2 (10 nM, 5 min perfusion) using increasing von Frey filaments (arrows) applied to the receptive field in the skin for a brief 5 s stimulus. Each point represents a single action potential. A , recoverable increase in number of action potentials and peak firing frequency after Cn2 application. B , A‐fibre with continuous burst firing after removal of stimulus. Inset, expanded view of characteristic spontaneous burst firing pattern, including volley of action potentials followed by period of quiescence. C , increase in number of action potentials of a subset of mechanically sensitive A‐fibres, those represented in panel A (action potentials at maximum von Frey stimulus, before: 81 ± 17 and after Cn2: 269 ± 65; P < 0.05, paired t test, n = 9). D , peak firing frequency of mechanically sensitive A‐fibres (panel A ) before (47.8 ± 16.5 Hz) and after Cn2 (210.8 ± 29.3 Hz) application (paired t test, P < 0.05, n = 12). E , conduction velocity is significantly different between the A‐fibres that display burst firing pattern such as panel B (3.5 ± 0.5 m/s; n = 5) and those which have increased response to mechanical stimulus such as panel A (7.3 ± 1.3 m/s; n = 12, Mann–Whitney test, P < 0.05). g, grams of stimulus.

Indeed, application of Cn2 at the distal terminals innervating the skin caused increased responses to mechanical stimulation in A‐fibres. Under control conditions, isolated mechanosensitive A‐fibres (both SA and RA) responded to mechanical stimuli of increasing force with a greater number of action potentials at higher frequency (Fig. 5 A ). The arrows (Fig. 5 A and B ) denote the brief 5 ms mechanical stimuli. After incubating the receptive fields of these fibres with Cn2 for 5 min, these A‐fibres became mechanically sensitized in two phenotypically distinct groups, henceforth termed non‐bursting and bursting. In the first (non‐bursting), the stimulus–response relationship was increased after Cn2 exposure, with each mechanical stimulus eliciting a greater number of action potentials at higher frequency than the corresponding control stimulus (Fig. 5 A , C and D ). Peak frequency of discharge was taken from the highest mechanical stimulus and was significantly increased compared to before Cn2 application (Fig. 5 D ). Correspondingly, the number of action potentials at this stimulus was significantly increased after Cn2 application (Fig. 5 C ). In the second group (bursting), a single brief (5 s) mechanical stimulus – which under control conditions elicited only a single discreet discharge of action potentials – led to sustained firing in a pattern of bursting action potential discharges that continued without any further mechanical stimulation for the duration of the recordings (up to 30 min) (Fig. 5 B , inset). Unlike other Na V activators such as the Na V 1.7 selective OD1 or P‐CTX‐1, Cn2 10 nM did not cause spontaneous firing in the absence of mechanical stimulus in any A‐fibre ( n = 23) (Deuis et al . 2016 ; Inserra et al . 2017 ). However, at higher concentrations (>100 nM) Cn2 induced spontaneous firing followed by loss of excitability. Non‐bursting fibres had faster conduction velocities than those with bursting phenotype (Fig. 5 E ). A‐fibre receptive fields were subject to cold and heat ramps and application of Cn2 did not lead to de novo sensitivity (data not shown).

Cultured DRG neurons are routinely used as a robust model of peripheral sensory neurons because they retain their endogenous ion channels necessary for action potential initiation and propagation. However, functional expression of ion channels, for example Na V 1.6 channels, at the distal terminals in the skin and viscera remains poorly understood. The distal terminals of primary afferents are the predicted sites of action potential generation in response to external stimuli including hot, cold and touch. To investigate the functional role of Na V 1.6 channels at peripheral terminals in the skin, we exploited Cn2 as a selective activator of Na V 1.6 channels to measure primary afferent response to mechanical stimulus in skin–saphenous nerve and colon–splanchnic nerve preparations. Given the dramatic effect on large‐diameter neurons, supported by the established expression pattern of Na V 1.6 channels, we hypothesized that Cn2 would alter firing of A‐fibres in the skin.

A , representative trace from current clamp recording of isolated large‐diameter DRG neurons before and after Cn2 (50 nM) application. Cn2 caused an increase in number of action potentials evoked by current injection at its respective current threshold (red). A decrease in evoked action potentials was observed following an injection of current 1.5 times its respective current threshold (blue). Current clamp protocol, inset. B , expanded view of the altered action potential shape shown in A . Arrows at interspike ramp point show change in appearance of second action potential.

Application of Cn2 onto large‐diameter DRG neurons caused a significant increase in the number of evoked action potentials at its respective current threshold (Fig. 4 A , red). Cn2 caused large‐diameter DRG neurons to transition from phasic to tonic firing. Increasing the current input to 1.5 times the current threshold reduced the number of action potentials elicited (Fig. 4 A , blue). The overall increase in activity was not associated with changes in input resistance, current threshold or action potential amplitude (Table 1 ). Interestingly, Cn2 altered the shape of the second action potential, making the interspike ramp steeper (Fig. 4 B ). By contrast, Cn2 did not induce repetitive firing patterns in Na V 1.6 −/− large‐diameter neurons (number of action potentials before and after Cn2 = 1, n = 8). Previously, native Cn2 was shown to significantly depolarize the membrane potential of Purkinje neurons (Schiavon et al . 2006 ), but we observed no significant difference in resting membrane potential of WT large‐diameter neurons during or after Cn2 application (Table 1 ). This is consistent with evidence that Na V 1.6 channels are critical for the fast upstroke of an action potential but do not contribute to the maintenance of the resting membrane potential (Royeck et al . 2008 ; Xie et al . 2013 ; Chen et al . 2018 ).

As toxins that enhance persistent (for example, δ‐AITX‐Avd1c from Anemonia sulcata (ATX‐II)) and resurgent current (Cn2) affect firing properties of cortical neurons (Mantegazza et al . 1998 ; Schiavon et al . 2006 ), we assessed the effect of Cn2 on excitability of large‐diameter DRG neurons. We found that control large‐diameter mouse DRG neurons have a high threshold and resisted repetitive firing under our culture conditions (Table 1 ).

A , representative trace of resurgent current from large DRG neurons in the presence and absence of 50 nM Cn2. DRG neurons were held at −100 mV and depolarized to +30 mV for 20 ms, followed by repolarization to a range of pulses (−10 to −80 mV) to elicit resurgent current. B , average resurgent current amplitude before and after Cn2 application shows a 3‐fold increase in peak amplitude (Control: 3.1 ± 0.7 nA; Cn2: 9.9 ± 1.0 nA; n = 7, P < 0.05), and partial reduction after a 5 min washout period. C , current–voltage relationship showing a hyperpolarized shift (∼40 mV) in peak resurgent current amplitude after Cn2 (50 nM), and the appearance of two peaks following a 5 min washout period (Control: −27.1 ± 3.2 mV; Cn2: −68.6 ± 2.1 mV; n = 7, P < 0.05). D , peak resurgent current measured before and after Cn2. E , peak voltage shifted ∼40 mV after Cn2 application.

The native Cn2 toxin enhances resurgent current in Purkinje neurons that exhibit high frequency firing and have high expression of Na V 1.6 channels (Schiavon et al . 2006 ). DRG neurons are also known to generate resurgent currents through Na V 1.6 (Cummins et al . 2005 ). Therefore, we asked whether Cn2 could enhance the endogenous resurgent current in large‐diameter DRG neurons. Treatment of DRG neurons with 50 nM Cn2 increased the endogenous resurgent current amplitude in large‐diameter neurons by 3‐fold (Fig. 3 A , B and D ). Cn2 (50 nM) dramatically shifted the voltage‐dependence of resurgent current approximately 40 mV in a hyperpolarizing direction (Fig. 3 C–E ). The increase in the amplitude of the resurgent current was partially reduced after 5 min of washout (Fig. 3 A , B and D ), and interestingly, the voltage‐dependence of the resurgent current shows two peaks, a depolarized peak at (−28.8 mV), which corresponds to the current in untreated cells, and the hyperpolarized peak (−67.9 mV), which corresponds to channels presumably still bound by Cn2 (Fig. 3 B , C and E ). Increased resurgent current amplitude and the shift in current–voltage relationship was not observed in buffer‐treated cells (average peak current amplitudes of resurgent current: Control: 3.3 ± 1.3 nA; Mock: 3.2 ± 1.1 nA; Wash: 3.1 ± 1.0 nA; peak current voltage: Control: −31 ± 3 mV; Mock: −32 ± 2 mV; Wash: −34 ± 3.0 mV; paired t test, P > 0.05, n = 4).

, normalized sodium current–voltage relationship of large‐diameter (>35 µm) DRG neurons before (black) and after (red) Cn2 (50 nM) perfusion. Cn2 causes early sodium channel opening (inset) (= 10)., protocol used to investigate voltage sensor trapping by Cn2 modified from (Schiavon)., representative trace (pulse to −50 mV) in the presence and absence of Cn2 (50 nM)., Cn2 significantly increases persistent current in large‐diameter DRG neurons (peak persistent at −35 mV: Control: 306.4 ± 70.6 pA; Cn2: 893.4 ± 224.1 pA;< 0.05,= 9).and, Cn2 does not alter current–voltage relationship of large‐diameter DRG neurons from Na1.6mice () or enhance persistent current (). Cn2 does not induce early opening in Na1.6large‐diameter neurons (, inset)., current–voltage relationship of WT small‐diameter neurons was not affected by Cn2 application., persistent current amplitude was not enhanced by Cn2 in small‐diameter DRG neurons.

To confirm that Cn2 is selective for Na V 1.6 channels in a native neuronal environment with heterogeneous and endogenous sodium channel expression, we utilized isolated DRG neurons from wild‐type (WT) and Na V 1.6 −/− ( SCN8A medtg/medtg ) mice. Since Cn2 preferentially activated large‐diameter DRG neurons (Fig. 1 ) in the Ca 2+ assay, these neurons (>35 µm) were initially chosen for voltage‐clamp recordings. In accord with previous findings, Cn2 (50 nM) caused sodium current influx at previously prohibitive potentials between −70 and −40 mV in these neurons (Fig. 2 A , inset). In control conditions, pulses to −50 mV elicit currents of an average amplitude of −270 ± 24 pA; after application of Cn2 pulses to −50 mV elicited significantly increased inward currents of an average amplitude of −2075 ± 429 pA ( P < 0.05, paired t test, n = 7). Concurrently, Cn2 caused a significant 4‐fold increase in Na V persistent current that peaked at –35 mV compared to control current (Control: −306 ± 77 pA; Cn2: −893.4 ± 224.1 pA; Fig. 2 C and D ). This enhanced persistent current was absent in both Na V 1.6 −/− large‐diameter neurons and WT small‐diameter neurons (Fig. 2 F–H ). Cn2 did not shift the V 1/2 activation or significantly alter the slope of large‐diameter neurons ( V 1/2 activation: Control: −25.6 ± 0.6 mV; Cn2: −25.2 ± 0.7 mV; slope: Control: 4.8 ± 0.5; Cn2: 5.8 ± 0.6; P > 0.05, paired t test, n = 10). Cn2 did not induce early opening in large‐diameter neurons from Na V 1.6 −/− mice confirming selective action at Na V 1.6 channels (Fig. 2 G , inset). Following, Cn2 did not shift the V 1/2 of activation or change the slope of activation in large‐diameter neurons from Na V 1.6 −/− mice ( V 1/2 activation: Control: −16.8 ± 0.8 mV; Cn2: −21.2 ± 1.1 mV; slope: Control: 6.7 ± 0.7; Cn2: 6.2 ± 1.0; paired t test, P > 0.05, n = 11). Furthermore, Cn2 (50 nM) failed to significantly alter the current–voltage relationship in any of the small‐diameter DRG neurons examined (Fig. 2 E , inset; small‐diameter DRG neuron V 1/2 activation: Control: −18.4 ± 1.1 mV; Cn2: 18.5 ± 0.9 mV; slope: Control: 8.8 ± 1; Cn2: 8.2 ± 0.8; paired t test, P > 0.05, n = 7).

A , DRG neurons before (background) and after addition of Cn2 (500 nM). Cn2 caused rapid, strong and sustained calcium influx in a subset of large‐diameter (white arrowheads) neurons. Small‐diameter neurons that responded with calcium influx are labelled with a red arrow. Scale bar is 100 µM. B , traces from all neurons (defined as cells responding to KCl (30 mM) and/or Cn2) of one representative experiment. C , percentage of large‐ (>600 µm 2 ), medium‐ (300–600 µm 2 ) and small‐diameter (<300 µm 2 ) neurons responsive to application of Cn2 (500 nM). Total number of neurons from 3 independent experiments is indicated. l.d., large diameter, m.d., medium diameter; s.d., small diameter.

Although the β‐scorpion toxin Cn2 was previously reported to display exquisite selectivity for Na V 1.6 channels over other Na V isoforms in HEK293 cells (Israel et al . 2018 ), neurons express a number of different voltage‐gated ion channel subtypes including those belonging to the K V , Ca V and Cl − families (Hille, 2001 ). Furthermore, it is well understood that auxiliary proteins such as sodium channel β‐subunits, calmodulin (CaM) and fibroblast growth factor homologous factors (FHFs) play an important role in modulating Na V 1.6 channel function (Herzog et al . 2003b ; Wittmack et al . 2004 ; Zhao et al . 2011 ). Within peripheral sensory neurons of DRGs, Na V 1.6 makes the largest contribution (50–60%) to the TTX‐S current in large and medium neurons. In addition, Na V 1.6 is also expressed in small‐diameter DRG neurons where it contributes roughly 30% of the TTX‐S current (Felts et al . 1997 ; Black et al . 2002 ; Chen et al . 2018 ). We utilized a high‐content calcium imaging assay to assess the effect of Cn2 on small‐, medium‐ and large‐diameter DRG neurons. Previously, Cn2 was shown to retain selectivity for Na V 1.6 channels in vitro up to 1 µM (Israel et al . 2018 ). Application of Cn2 (500 nM) to dissociated mouse DRG neurons caused an immediate sharp and sustained increase in intracellular calcium concentration ([Ca 2+ ] i ) in a large proportion (45%) of large‐diameter neurons (Fig. 1 ). A small population of medium‐ and small‐diameter neurons also responded to Cn2, consistent with the reported expression profile of Na V 1.6 channels (Felts et al . 1997 ; Black et al . 2002 ; Chen et al . 2018 ).

Discussion

The expression of different Na V subtypes in distinct populations of peripheral sensory neurons is inextricably linked to the diverse function of these neurons. The largely peripherally restricted Na V channel subtypes Na V 1.7, Na V 1.8 and Na V 1.9 have long been implicated in various human pain conditions, including peripheral neuropathy and congenital insensitivity to pain (Dib‐Hajj et al. 2013, 2015; Erickson et al. 2018). A contribution of Na V 1.6 channels to enhanced excitability of sensory neurons is supported by evidence from human genetic studies and murine models suggesting that Na V 1.6 plays a significant role in the development and maintenance of trigeminal neuralgia and neuropathic pain (Ren et al. 2012; Xie et al. 2013; Tanaka et al. 2016; Chen et al. 2018). This emerging evidence supports a role for Na V 1.6 in pain, but additional insight into its specific role in nociceptive signalling is needed.

In the present study, we unequivocally confirmed the selectivity of the β‐scorpion toxin Cn2 as a selective activator of Na V 1.6 channels using DRG neurons isolated from WT and Na V 1.6−/− (SCN8Amedtg/medtg) mice. We showed an increase in excitability of large‐diameter sensory neurons in both in vitro cultures and ex vivo skin–nerve preparations. Selective Na V 1.6 activation caused striking sensitivity to mechanical stimulation due to action in myelinated A‐fibres in the skin. Consistent with expression of Na V 1.6 along unmyelinated fibres (Black et al. 2002), we found that Cn2 is able to increase the response of colonic nociceptors (C‐fibres) to mechanical stimulation. Taken together, our data support an essential role for Na V 1.6 in peripheral sensory neurons, specifically at the distal terminals of mechanosensing fibres innervating the skin and colon.

Cn2 is a canonical β‐scorpion toxin, previously shown to induce current at prohibitively hyperpolarized potentials in HEK293 cells expressing Na V 1.6 channels (Schiavon et al. 2006). In the current study, we confirmed specificity of this toxin to Nav1.6 and toxin‐induced early activation of Na V 1.6 channels in native sensory neurons. Additionally, Cn2 application on DRG neurons led to a robust persistent current not previously reported in the HEK293‐Na V 1.6 studies (Schiavon et al. 2006; Israel et al. 2018). This is probably due to the different cellular background in these studies and differential expression of key interacting proteins including Na V β subunits, which are known to regulate persistent sodium currents (such as β4) (Aman et al. 2009; Bant & Raman, 2010).

Specific deletion of Na V 1.6 channels from Na V 1.8‐expressing small‐diameter sensory neurons decreased current density by approximately 30% (Chen et al. 2018). However, Na V 1.6 channel activation by Cn2 did not alter sodium currents in small‐diameter DRG neurons, perhaps because it is masked by the predominant Nav1.7 TTX‐S current in these neurons. As cells were not selected based on evidence of expression of Na V 1.6 channels in this study and a small but distinct number of small‐diameter neurons responded to Cn2 in high‐throughput calcium imaging, we cannot rule out a functional contribution of Na V 1.6 channels to a subset of small‐diameter DRG neurons. Enhanced resurgent currents in sensory neurons are thought to contribute to various pain disorders including paroxysmal extreme pain disorder and oxaliplatin‐induced cold allodynia (Jarecki et al. 2010; Sittl et al. 2012). Consistent with findings from mouse Purkinje neurons, which express high levels of Na V 1.6 and respond to Cn2 with accelerated firing and gradual spike accommodation during depolarizing current injection (Schiavon et al. 2006), Cn2 treatment caused an increase in resurgent current in large‐diameter DRG neurons and enhanced repetitive firing of these neurons, which was followed by gradual spike accommodation.

The Cn2‐induced increase in the persistent and resurgent currents in large DRG neurons could be caused by Na V β subunits, which are known to regulate these currents, for example β4 subunits (Aman et al. 2009; Bant & Raman, 2010). Na V β4 is highly expressed in neurofilament‐positive neurons that correspond to heavily myelinated, large‐diameter A‐fibres: those affected by Cn2 in this study (Lawson & Waddell, 1991; Usoskin et al. 2015). The presence of a β‐scorpion toxin, such as Cn2, can trap the DIIS4 segment in the outward position and delay channel closing (Cestele et al. 1998; Schiavon et al. 2006). This could make more of the channels available for the open channel blocker, for example the C‐terminal tail of the β4 subunit (Grieco et al. 2005). Thus, the 3‐fold increase in resurgent current might be due at least partially to a cumulative action with the β4 subunit in large‐diameter sensory neurons.

Although a hyperpolarized shift in Na V activation was absent in large‐diameter DRG neurons from Na V 1.6−/− mice, it is possible that Cn2, like other animal‐derived toxins, may modulate other ion channels that alter firing properties in neurons, such as voltage‐gated potassium channels (Chi & Nicol, 2007; Redaelli et al. 2010; Liu & Bean, 2014). However, Cn2 did not alter firing patterns of DRG neurons from Na V 1.6−/− mice, confirming that the distinct firing pattern is driven by Na V 1.6 channel activation alone. Furthermore, unlike modulators of voltage‐gated potassium channels, Cn2 did not change the resting membrane potential, rheobase, after‐hyperpolarization amplitude or action potential width in wild‐type DRG neurons. Although the opening at hyperpolarized potentials caused by Cn2 is the canonical β‐scorpion effect, it remains small compared to the substantial increase in persistent current, peak resurgent current and the strong hyperpolarizing shift in resurgent current. Therefore, we posit that the Cn2‐induced changes in resurgent current amplitude and voltage‐dependence seen in large‐diameter DRG neurons (>50% increase, 40 mV hyperpolarizing shift) underlie the repetitive firing induced in large‐diameter sensory neurons.

Sensory neural transduction begins at the distal terminals in the skin, where a range of ion channels and transducer channels detect external stimuli. Evidence from immunostaining of skin preparation confirms the presence of Na V 1.6 channels in these terminals in the skin (Persson et al. 2010). Cn2 altered the excitability of large‐diameter DRG soma that give rise to myelinated Aδ‐ and Aβ‐fibre projections into the skin. Accordingly, Cn2 preferentially enhanced activity of these fibre types ex vivo. Na V 1.6 activation by Cn2 also causes robust mechanical allodynia in vivo. This enhanced response to mechanical stimulus is mirrored ex vivo in the skin in a subset of A‐fibres that have conduction velocities in the Aδ and Aβ range (Harper & Lawson, 1985; Zimmermann et al. 2009). Exaggerated responses to non‐painful mechanical stimulus in vivo may arise from the increased action potential number and frequency at relatively innocuous (e.g. 0.5 grams force) stimulus observed in ex vivo recordings.

Immunocytochemistry demonstrates robust Na V 1.6 expression along non‐myelinated PNS axons, corresponding to C‐fibres within the epidermis (Black et al. 2002). However, some ex vivo and in vivo findings from this study suggest a minor functional role of Na V 1.6 in these neurons in the skin, which may be due to differential expression of sodium channel isoforms at the C‐fibre distal segment, or to the mismatch between the representation of Na V 1.6 in the plasma membrane of the terminals versus the cytoplasmic pool, which cannot be distinguished using immunostaining of permeabilized tissue and antibodies that only recognize intracellular epitopes as is the case with the antibodies that were used in these studies. A higher density of Na V 1.6 channels within terminals in a subpopulation of C‐fibres in the skin might explain the subtle changes (to temperature and mechanical stimulus) observed in the study. This aspect could be explored with the use of selective molecules in conjunction with knock‐out animals.

Intriguingly, our data suggest the Na V 1.6 isoform is also important for mechanical excitability of peripheral neurons innervating the gut. Visceral afferents are solely composed of small‐diameter thinly myelinated Aδ‐ and non‐myelinated C‐fibres (Brierley et al. 2011; Brierley & Linden, 2014; Erickson et al. 2018; Sadeghi et al. 2018). However, single‐cell RT‐PCR data show that 62.5% of colon‐innervating thoracolumbar DRG neurons express Na V 1.6 channels (Inserra et al. 2017). Indeed, selective antagonism of Na V 1.6 channels in low‐threshold colorectal afferents reduced tonic spiking in response to stretch (Feng et al. 2015). Our evidence supports this finding, extends it to high‐threshold nociceptors, and suggests that mechanosensitive C‐fibres in the gut express Na V 1.6 channels and activation of this isoform leads to increased mechanosensitivity. Surprisingly, while Cn2 did not change the voltage–current relationship in small‐diameter DRG neurons in culture, Na V 1.6 activation in the skin led to a small but significant reduction in cold‐induced action potential firing in a subset of C‐fibres. Consistent with this unexpected decrease in excitability during cooling, Cn2 alone does not lead to sensitization to cold stimulus in vivo. While the molecular mechanisms of this effect are not known, it is conceivable that cooling‐induced slow inactivation develops more readily in the presence of Cn2. Nevertheless, this observation clearly supports that Na V 1.6 is indeed expressed in unmyelinated C‐fibres, albeit the precise functional role of this isoform in these and other subtypes of small‐diameter unmyelinated fibres remains unresolved.

The inability of Cn2 to induce spontaneous firing in A‐fibres at concentrations that induced spontaneous pain in vivo might be explained by the unique β‐scorpion toxin mechanism of action (Cestele et al. 1998). In the model, the Na V 1.6 voltage sensor must move outward, which requires an initial depolarization, allowing toxin binding that ultimately facilitates early channel opening. Indeed, without this depolarizing step, other β‐toxins cause only a decrease in peak current (Cestele et al. 1998; Leipold et al. 2012). It is plausible in this experimental design that this initial depolarizing step ex vivo may be arrived at by mechanical stimulation with von Frey hair – or indeed in vivo by standing. Mechanotransducer channels expressed at the distal terminal depolarize the membrane allowing Cn2 binding to Na V 1.6 channels, and subsequent action potentials are then affected by the Cn2–Na V 1.6 complex. Current clamp experiments from sensory neurons (this study) and Purkinje neurons (Schiavon et al. 2006) also support this hypothesis. Characteristics of the first action potential remain unchanged while the increased resurgent current promotes a steeper interspike ramp of subsequent action potentials. Resurgent current facilitates repetitive and sustained action potential firing (Raman & Bean, 1997; Bant & Raman, 2010), but Cn2‐induced repetitive firing undergoes adaptation and decrease in amplitude which might be due to cumulative sodium channel inactivation and absence of supporting potassium currents for sustained firing. Further examination with peptide analogues that have a reduced capacity for voltage‐sensor trapping, such as the Cn2‐E15R mutant, may help elucidate this mechanism (Karbat et al. 2010; Israel et al. 2018).