Contribution of tetrodotoxin-resistant persistent Na⁺ currents to the excitability of C-type dural afferent neurons in rats

Neurons within the TG innervating the dura were identified after the application of the retrograde tracer DiI to the dura, as previously described [23, 24]. Briefly, male Sprague Dawley rats (three to four weeks old; Samtako, Osan, Republic of Korea) were intraperitoneally anesthetized with a mixture of ketamine (20 mg/kg) and xylazine (10 mg/kg). The cranial bone overlying the superior sagittal sinus was gently removed using a careful craniotomy procedure involving a dental drill, and the dura was exposed with a burr hole of 2 mm. Ten microliters of DiI solution, in which 100 mg/ml of DiI in dimethyl sulfoxide (DMSO) was diluted 1:10 (v/v) with saline, was applied to the dura. One minute after the application of DiI solution, a dental resin was placed on the exposed dura to replace the removed cranial bone. The incision was closed with sutures, and the rats received intramuscular injections of penicillin G (100,000 U/kg) and naproxen (10 mg/kg) to reduce the risk of postoperative infection and pain. At 7–10 days after the DiI application, the rats were decapitated under ketamine anesthesia (100 mg/kg administered intraperitoneally). A pair of TG were dissected and treated with an external solution [in mM: 150 NaCl, 3 KCl, 2 CaCl₂, 1 MgCl₂, 10 glucose and, 10 Hepes (pH 7.4 with Tris-base)] containing 0.3% collagenase (type I) and 0.3% trypsin (type I) for 40–60 min at 37 °C. Thereafter, the TG neurons were dissociated mechanically using trituration with fire-polished Pasteur pipettes in a culture dish (Primaria 3801; Becton Dickinson, Rutherford, NJ, USA). The isolated neurons were used for electrophysiological recordings 2–6 h after preparation. Images of the TG neurons were obtained using a digital microscope camera (ProgRes® MF; Jenoptik, Jena, Germany). For the other subset experiments, TG neurons were obtained from adult male or female rats (8–9 weeks old; Samtako).

Previous studies have shown that acute bath application of a mixture of IMs (1 μM PGE₂, 10 μM bradykinin, and 1 μM histamine) sensitizes dural afferent neurons and increases their excitability [23, 25, 26]. In addition, bath application of a mixture of IMs increased the peak amplitude of TTX-R I_Na [26]. However, we examined whether peripheral sensitization by the application of IMs to the dura mater affected TTX-R I_Na, including transient and persistent currents and the excitability of dural afferent neurons. Since the identification of dural afferent neurons by the retrograde fluorescent dye DiI takes over 7 days, we applied a mixture of IMs to dural afferent neurons with chronic treatment methods using Elvax (ethylene–vinyl acetate copolymer resin) implantation incorporated with IMs. Although chronic treatment with IMs using Elvax implantation to the dura mater has not been described, Elvax implantation has been widely used for sustained release of small molecules, such as TTX [27], and peptides or proteins [28, 29].

To prepare Elvax implants, beads (100 mg) of Elvax (40 W; DuPont; kindly gifted from JS POLYMER, Seongnam, Republic of Korea) were dissolved in dichloromethane (100 mg/ml) and homogeneously mixed with 20 μl of DMSO containing IMs, i.e., 10 mM PGE₂, 20 mM bradykinin, and 10 mM histamine. Thereafter, the Elvax solution was dropped on a glass dish using a 1-ml syringe, rapidly frozen, kept at -80 °C for 1 h, and maintained at -20 °C overnight to allow the dichloromethane to evaporate. To implant an Elvax piece containing IMs, male Sprague Dawley rats (3–4 weeks old; Samtako) were intraperitoneally anesthetized with a mixture of ketamine (20 mg/kg) and xylazine (10 mg/kg). Two burr holes (2 mm diameter) were made using a careful craniotomy procedure, and the dura was exposed. DiI solution and a small Elvax piece (approximately 2 × 2 × 2 mm, 8.0 ± 0.3 mg, n = 11) were placed on the dura mater through each hole in the cranial bone, and a dental resin was placed on the exposed dura to replace the removed cranial bone. The Elvax piece was usually left on the dura mater from the day of implantation until the day of electrophysiological recording (7–10 days). The implantation of Elvax containing vehicle (DMSO) or IMs did not cause deficits in the general motor behaviors of rats. Considering that IMs were equally distributed in Elvax, the amounts of PGE₂, bradykinin, and histamine in the implanted Elvax piece (8 mg) were approximately calculated to be 2.9, 340, and 5.6 μg, respectively. In the present study, we did not measure the total estimated concentration of IMs in the dural tissue from the Elvax piece, the duration of IM release after the implantation of Elvax, and the residual amounts of IMs within the implanted Elvax piece after the animals were euthanized. However, biomolecules or chemicals (< 2 kDa), such as basic fibroblast growth factor, MK-801, and doxycycline, embedded in the Elvax, have been known to be released over 7 days with a peak on the first day [30‐32]. In the present study, since we did not perform any behavioral test to examine migraine-like pain behaviors for the Elvax-IM implantation in vivo, we could not verify whether the Elvax-IM implantation in the dura mater induces migraine-like pain behaviors. Further behavioral studies are needed to validate the Elvax-IM implantation as a chronic inflammation model for peripheral sensitization of the dura mater.

All electrical measurements were performed using conventional whole-cell patch recordings and a standard patch-clamp amplifier (Axopatch 200B; Molecular Devices, Union City, CA, USA). Neurons were voltage-clamped at a holding potential (V_H) of -80 mV, except where indicated. Patch pipettes were produced from borosilicate capillary glass (G-1.5; Narishige, Tokyo, Japan) using a pipette puller (P-97; Sutter Instrument Co., Novato, CA, USA). The resistance of the recording pipettes filled with the internal solution was 0.7–1.2 MΩ. Membrane potentials were corrected for the liquid junction potential, and the pipette capacitance and series resistance (60%–90%) were compensated for. DiI-positive TG neurons were viewed under phase contrast or fluorescence using an inverted microscope (TE2000; Nikon, Tokyo, Japan). Membrane currents were filtered at 3–5 kHz, digitized at 10–20 kHz, and stored on a computer equipped with pCLAMP 10.7 (Molecular Devices). Capacitative and leak currents were subtracted using the P/4 subtraction protocol of pCLAMP 10.7. To record the TTX-R I_Na, the internal solution was composed of the following (in mM): 140 CsF, 10 CsCl, 2 EGTA, 2 Na₂-ATP, and 10 Hepes (pH 7.2 with Tris-base). In the current-clamp experiments, both CsF and CsCl were replaced with equimolar KF and KCl, respectively. All experiments were performed at room temperature (22 ºC–25 ºC). The bath solution used was composed of the following (in mm): 130 NaCl, 20 TEA-Cl, 2 CaCl₂, 1 MgCl₂, 10 Hepes, 10 glucose, 0.01 CdCl₂, and 0.0003 TTX (pH 7.4 with Tris-base).

The peak amplitude of transient TTX-R I_Na was measured by subtracting the baseline from the peak amplitude of TTX-R I_Na using pCLAMP 10.7. The amplitude of the steady-state component of transient TTX-R I_Na was measured by subtracting the baseline from the mean amplitude of TTX-R I_Na at 90–95 ms. For experiments in which the voltage-activation relationship was assessed, the extracellular Na⁺ concentration was reduced to 20 mM by replacing 110 mM NaCl with equimolar N-methyl-D-glucamine-Cl, and the amplitude of TTX-R I_Na was transformed into conductance (G) using the following equation:

G = I / (V—E_Na),

where E_Na is the Na⁺ equilibrium potential calculated using the Nernst equation. The voltage-activation and voltage-inactivation relationships of TTX-R Na⁺ channels were fitted to Boltzmann equations, respectively, as follows:

G/G_max = 1/{1 + exp[(V_{50,activation}—V)/k]} and I/I_max = 1—1/{1 + exp[(V_{50,inactivation}—V)/k]},

where G_max and I_max are the maximum conductance and current amplitude, respectively, V_{50,activation} and V_{50,inactivation} are half-maximal voltages for activation and fast inactivation, respectively, and k is the slope factor. The kinetic data for the recovery from inactivation were best fitted to the triple exponential function using the following equation:

I(t) = A₀ + A_fast × [1—exp(-t/τ_fast)] + A_intermediate × [1—exp(-t/τ_intermediate)] + A_slow × [1—exp(-t/τ_slow)],

where I(t) is the amplitude of TTX-R I_Na at time t, and A_fast, A_intermediate, and A_slow are the amplitude fraction of τ_fast, τ_intermediate, and τ_slow, respectively. The kinetic data for the development of inactivation were best fitted to the double exponential function using the following equation:

I(t) = A₀ + A_fast × [exp(-t/τ_fast)] + A_slow × [exp(-t/τ_slow)],

where I(t) is the amplitude of TTX-R I_Na at time t, and A_fast and A_slow are the amplitude fraction of _fast and _slow, respectively. Numerical values are provided as the mean ± standard error of the mean (SEM) using values normalized to the control. Significant differences were tested using unpaired t-tests, except where indicated. P values of < 0.05 were considered statistically significant. In the current-clamp experiments, the rheobase currents, which represent the minimal currents that generate action potentials, were determined using a square depolarizing current injection in 10 pA increments (500 ms duration).

The following drugs were used in this study: riluzole, collagenase, trypsin, TTX, DiI, prostaglandin E₂, bradykinin, histamine (all from Sigma, St. Louis, MO, USA), A803467, and ZD7288 (the latter two from Tocris, Bristol, England). ZD7288 and TTX were dissolved in distilled water to give a stock solution of 50 mM and 3 mM, respectively. Riluzole and A803467 were dissolved in DMSO to give a stock solution of 100 mM and 1 mM, respectively. The final concentration of DMSO applied to the external solution was ≤ 0.1% v/v, and DMSO (0.1% v/v) did not affect the TTX-R I_Na (Supplementary Fig. S1A). Extracellular solutions containing the drugs, except collagenase, trypsin, and DiI, were applied using the “Y-tube system” for rapid solution exchange [33].

The voltage-activation relationship of TTX-R Na⁺ channels was examined in small- and medium-sized DiI-positive neurons. In these experiments, extracellular Na⁺ concentration was reduced to 30 mM to improve the quality of voltage clamping. TTX-R I_Na were recorded using 100 ms depolarizing test pulses from -80 mV to + 20 mV in 10 mV increments (Supplementary Fig. S2A). The conductance data were then normalized to the maximal conductance and fitted to the Boltzmann function (Supplementary Fig. S2A). However, the midpoint voltages for activation (V_{50,activation}) did not differ between the two groups of DiI-positive neurons [V_{50,activation}: -22.8 ± 0.3 mV (n = 32) and -22.3 ± 0.3 mV (n = 44) in small- and medium-sized DiI-positive neurons, respectively; p = 0.25]. In addition, the V_{50,activation} values were not related to cell capacitance (r = 0.01, n = 76, p = 0.95; Supplementary Fig. S2B) or the density of TTX-R I_NaP (r = 0.02, n = 76, p = 0.85; Supplementary Fig. S2C). Contrastingly, Na_V1.9-mediated TTX-R I_Na were clearly observed at a V_H of -120 mV, where the current density of Na_V1.9-mediated TTX-R I_Na did not differ between the small- and medium-sized DiI-positive neurons (Supplementary Fig. S2D, E).

The steady-state fast inactivation of TTX-R Na⁺ channels was also examined in small- and medium-sized DiI-positive neurons. TTX-R I_Na were recorded using 100 ms depolarizing test pulses to -10 mV following 500 ms prepulses from -120 to -10 mV in 10 mV increments (Fig. 3A, Ba). The peak currents were then normalized to the maximal current recorded at a prepulse of -120 mV, and the data were fitted to the Boltzmann function (Fig. 3Bb). The midpoint voltages for inactivation (V_{50,inactivation}) in small-sized neurons were significantly lower than those in medium-sized neurons, where the V_{50,inactivation} values were correlated with cell capacitance (r = 0.64, p < 0.01; Fig. 3Ca). The mean V_{50,inactivation} values were -32.6 ± 0.7 mV (n = 32) and -26.9 ± 0.4 mV (n = 44) in small- and medium-sized DiI-positive neurons, respectively (p < 0.01; Fig. 3Cb). The V_{50,inactivation} values were further correlated with the density of TTX-R I_NaP (r = 0.86, p < 0.01; Fig. 3D). We found that the values of V_{50,inactivation} were more negative in small-sized DiI-positive neurons than in their medium-sized counterparts, which may have been due to the faster shift in V_{50,inactivation} during the recording, given that F⁻ is known to elicit a hyperpolarizing shift of V_{50,inactivation} values. However, in contrast to Na_V1.9, there is evidence that Na_V1.8 is not modulated by F⁻ [42]. Nevertheless, we found a similar time-dependent negative shift in V_{50,inactivation} values for the two types of DiI-positive neurons (Fig. 3E), indicating that a hyperpolarizing shift in V_{50,inactivation} during recording might not be responsible for the difference in V_{50,inactivation} values observed between small- and medium-sized neurons. Therefore, we further examined the effects of prepulse duration on steady-state fast inactivation. The V_{50,inactivation} values were found to be related to prepulse duration in the two DiI-positive neuron types. The voltage dependence of fast inactivation did not differ under prepulse duration conditions of ≤ 30 ms (Fig. 3F). We also found no difference between the V_{50,inactivation} values obtained from TTX-R I_NaT and TTX-R I_NaP in medium-sized DiI-positive neurons (Supplementary Fig. S1F, G), indicating that both TTX-R I_NaT and TTX-R I_NaP are mediated by the same channels.

Our finding that the voltage-fast inactivation relationships for TTX-R Na⁺ channels depend on prepulse duration suggests that slow inactivation might underlie the observed difference in V_{50,inactivation} values between the two DiI-positive neuron types. Therefore, we further examined the voltage dependence of slow inactivation of TTX-R Na⁺ channels in small- and medium-sized DiI-positive neurons. TTX-R I_Na were elicited using 100 ms depolarizing test pulses to -10 mV following 5,000 ms prepulses from -100 to -10 mV in 10-mV increments (Fig. 3G, Ha). Peak currents were normalized to the maximal current recorded at a prepulse of -100 mV, and the data were fitted to the Boltzmann function (Fig. 3Hb). The V_{50,inactivation} values for slow inactivation were significantly lower in small-sized neurons than those in medium-sized neurons, where V_{50,inactivation} was -49.2 ± 0.8 mV (n = 30) and -41.1 ± 0.4 mV (n = 41), respectively (p < 0.01). The V_{50,inactivation} values were also correlated with cell capacitance (r = 0.69, n = 71, p < 0.01; Fig. 3Ia) and the density of TTX-R I_NaP (r = 0.81, n = 71, p < 0.01; Fig. 3Ib).

Both the inactivation and recovery kinetics of voltage-gated Na⁺ channels directly influence the availability of Na⁺ channels during repetitive action potentials. Therefore, the slow inactivation kinetics of TTX-R Na⁺ channels were examined using a two-pulse protocol in small- and medium-sized DiI-positive neurons (Supplementary Fig. S3A; Fig. 4Aa). The amplitude ratios of two TTX-R I_Na (P₂/P₁) were calculated and plotted against the prepulse duration for inactivation (Fig. 4Ab). The P₂/P₁ ratios were well-fitted to the double exponential function, which resulted in fast and slow time constants (τ_fast and τ_slow) (Fig. 4Ab), suggesting that TTX-R Na⁺ channels fall into the slow inactivation category with two distinct forms of kinetics, i.e., fast and slow kinetics. There were large differences between the inactivation kinetics of small- and medium-sized DiI-positive neurons (Fig. 4Ab). τ_fast and τ_slow values were correlated with neuronal size (τ_fast: r = 0.57; τ_slow: r = 0.41; n = 41; p < 0.01; Fig. 4Ba and Supplementary Fig. S3C). The mean τ_fast values were 352.1 ± 55.3 ms (n = 28) and 887.8 ± 52.9 ms (n = 47) for small- and medium-sized DiI-positive neurons, respectively (p < 0.01; Fig. 4Bb). The mean τ_slow values were also significantly smaller in small-sized DiI-positive neurons than those in medium-sized neurons [5.6 ± 0.7 s (n = 28) vs. 9.9 ± 0.7 s (n = 47), respectively; p < 0.01; Supplementary Fig. S3B]. However, the amplitude fractions of τ_fast and τ_slow (A_fast and A_slow, respectively) did not differ between the two DiI-positive neuron types (Supplementary Fig. S3B). Since the amplitude fraction of τ_fast was much larger than that of τ_slow (Supplementary Fig. S3B), the overall inactivation kinetics of TTX-R Na⁺ channels might be determined by τ_fast in both DiI-positive neuron types. τ_fast was further correlated with the density of TTX-R I_NaP in DiI-positive neurons (r = 0.69, n = 75, p < 0.01; Fig. 4C). However, A_fast and A_slow were not correlated with neuronal size or the density of TTX-R I_NaP (Supplementary Fig. S3C, D).

The extent of slow inactivation when using the two-pulse protocol depends on the duration of recovery time after the inactivation prepulse. Therefore, we examined whether inactivation kinetics were affected by changing the recovery time duration (Fig. 4D). Although the inactivation kinetics differed considerably between small- and medium-sized DiI-positive neurons (Fig. 4E), τ_fast values were not related to the recovery time duration in the two neuron types (Fig. 4E, F). However, A_fast values decreased as the recovery time duration increased (Fig. 4E, F). These results suggest that the recovery time duration after the inactivation prepulse might determine the extent rather than the kinetics of the fast component of slow inactivation. We also examined the effects of holding potentials on the inactivation kinetics of TTX-R Na⁺ channels. The two-pulse protocol was again applied to small- and medium-sized DiI-positive neurons at three different holding potentials (-60, -80, and -100 mV) (Fig. 4G). τ_fast was not dependent on the holding potentials in small- and medium-sized DiI-positive neurons. However, the A_fast values increased as the holding potentials increased (Fig. 4H, I). Thus, neither the recovery time duration nor holding potentials are responsible for the marked difference in the fast component of slow inactivation between small- and medium-sized DiI-positive neurons.

The extent of recovery from inactivation of TTX-R Na⁺ channels was also determined using the two-pulse protocol in small- and medium-sized DiI-positive neurons (Fig. 5A; Supplementary Fig. S4A). P₂/P₁ ratios were plotted against the prepulse duration for recovery times and fitted to the triple exponential function, which resulted in fast, intermediate, and slow time constants (τ_fast, τ_intermediate, and τ_slow, respectively) (Fig. 5A), suggesting that TTX-R Na⁺ channels recover from inactivation with three distinct forms of kinetics, i.e., fast, intermediate, and slow recovery kinetics. The recovery kinetics of small- and medium-sized DiI-positive neurons also differed substantially (Fig. 5Ab). τ_fast values were correlated with the density of TTX-R I_NaP (r = 0.75, n = 51, p < 0.01; Fig. 5Ba) but τ_intermediate and τ_slow values were not (τ_intermediate: r = 0.04; τ_slow: r = 0.19; n = 51; Supplementary Fig. S3C). The mean τ_fast values were 8.4 ± 0.9 ms (n = 18) and 1.9 ± 0.2 ms (n = 33) for small- and medium-sized DiI-positive neurons, respectively (p < 0.01; Fig. 5Bb). However, the mean τ_intermediate and τ_slow values did not differ between the two neuron types (Supplementary Fig. S4B). In contrast to the inactivation kinetics, the amplitude fractions differed substantially for τ_fast, τ_intermediate, and τ_slow (A_fast, A_intermediate, and A_slow, respectively) (Supplementary Fig. S4B). Taken together, the fast component of recovery from inactivation likely determines the overall recovery kinetics of TTX-R Na⁺ channels between small- and medium-sized DiI-positive neurons. τ_fast values were correlated with the density of TTX-R I_NaP (r = 0.75, n = 51, p < 0.01; Fig. 5C) but τ_intermediate and τ_slow values were not (Supplementary Fig. S4D). A_fast, A_intermediate, and A_slow were further correlated with neuronal size and the density of TTX-R I_NaP (Supplementary Fig. S4C, D).

Because recovery from inactivation should be related to the extent of inactivation, we examined whether recovery kinetics were affected by the duration of the conditioning prepulses used for channel inactivation (Fig. 5D). The extent of recovery from inactivation was greatly accelerated by shortening the inactivating prepulse duration but slowed down by lengthening this duration in small-sized, and to a lesser extent, medium-sized DiI-positive neurons (Fig. 5E). In particular, when TTX-R Na⁺ channels were inactivated with a prepulse duration of 50 or 5,000 ms, there was no difference in τ_fast values between the two DiI-positive neuron types [50 ms prepulse duration: 1.0 ± 0.2 ms (n = 9) and 0.8 ± 0.1 ms (n = 7) for small- and medium-sized neurons, respectively (p = 0.11); 5,000 ms prepulse duration: 11.2 ± 1.9 ms (n = 8) and 6.6 ± 1.7 ms (n = 8), respectively (p = 0.10); Fig. 5Fa]. The A_fast values were also increased by shortening the inactivating prepulse duration in small- and medium-sized DiI-positive neurons (Fig. 5Fb). However, when the TTX-R Na⁺ channels were inactivated with a prepulse duration of 50 or 5000 ms, there was no difference in the A_fast values between the two neuron types [50 ms prepulse duration: 0.80 ± 0.07 ms (n = 9) and 0.89 ± 0.05 (n = 7) for small- and medium-sized DiI-positive neurons, respectively (p = 0.28); 5,000 ms prepulse duration: 0.06 ± 0.03 (n = 8) and 0.12 ± 0.04 ms (n = 8), respectively (p = 0.09); Fig. 5Fb].

The use dependence of TTX-R Na⁺ channels was also examined in small- and medium-sized DiI-positive neurons. TTX-R I_Na were recorded using 20 successive depolarizing test pulses (30 ms duration) from -80 mV to -10 mV every 200 ms (5 Hz) (Supplementary Fig. S5Aa). The peak amplitudes of TTX-R I_Na were then normalized to that elicited by the first test pulse (Supplementary Fig. S5Ab). The extent of use-dependent inactivation was larger in small-sized DiI-positive neurons than that in medium-sized neurons, where the P₂₀/P₁ ratio of TTX-R I_Na was 0.85 ± 0.01 (n = 32) and 0.94 ± 0.1 mV (n = 44), respectively (p < 0.01; Supplementary Fig. S5Bb). The P₂₀/P₁ ratio of TTX-R I_Na was correlated with cell capacitance (r = 0.62, n = 74, p < 0.01; Supplementary Fig. S5Ba) and the density of TTX-R I_NaP (r = 0.72, n = 74, p < 0.01; Supplementary Fig. S5C).

The above-reported results suggest that the density of TTX-R I_NaP might be causally related to the extent of channel inactivation during sustained membrane depolarization. Therefore, we examined whether TTX-R I_NaP contribute to the excitability of dural afferent neurons under current-clamp conditions. The basal membrane properties, such as resting membrane potentials and rheobase currents, which are the minimal threshold currents that trigger action potentials, are summarized in Supplementary Table S1. When the integers of threshold currents (1 T–4 T) were applied to DiI-positive neurons, phasic and tonic firing patterns were detected in most small- and medium-sized DiI-positive neurons, respectively (Fig. 6A; Supplementary Table S1), which is consistent with our previous study [24]. In the presence of TTX (300 nM), the number of action potentials triggered by depolarizing current stimuli was slightly but significantly decreased in both neuron types (Fig. 6A, B). In addition, small- and medium-sized DiI-positive neurons still exhibited phasic and tonic firing patterns, respectively, even in the presence of TTX (Fig. 6A, B). We also examined the effects of riluzole treatment on the excitability of these neurons. Riluzole treatment was found to significantly reduce the number of action potentials elicited by depolarizing current stimuli in a concentration-dependent manner. Specifically, 10 μM riluzole significantly reduced the number of action potentials elicited by 4 T stimulation in small-sized [6.0 ± 0.7 and 1.3 ± 0.2 under control and riluzole conditions, respectively (n = 6); p < 0.01] and medium-sized (46.3 ± 7.7 and 3.8 ± 0.8 under control and riluzole conditions, respectively (n = 6); p < 0.01] DiI-positive neurons (Fig. 6C, D). Hyperpolarization-activated and cyclic nucleotide-gated (HCN) channels might contribute to the tonic firing pattern observed in medium-sized DiI-positive neurons because these channels are known to contribute to neuronal excitability with regular firing properties in various central neurons [43]. However, riluzole had no inhibitory effect on the sag potentials, which were greatly decreased by the HCN channel blocker ZD7288 (50 μM) (Fig. 6Cb).

However, riluzole at micromolar concentrations is known to activate small-conductance Ca²⁺-activated K⁺ (SK_Ca) channels [34]. Since SK_Ca channels contribute to afterhyperpolarization (AHP) of the action potential [44], riluzole might affect the activity of SK_Ca channels to reduce the frequency of action potentials in response to depolarizing current injection. Therefore, we examined whether riluzole affects AHP of action potential in DiI-positive neurons. Figure 6E shows the single action potentials elicited by brief depolarizing current stimuli (3 ms duration) in the absence and presence of 10 μM riluzole in medium-sized DiI-positive neurons. Riluzole (10 μM) had no effect on the amplitude of AHP (98.3 ± 0.8% of the control, n = 6, p = 0.08) and decay time constant of AHP (98.4 ± 4.3% of the control, n = 6, p = 0.61, Fig. 6E, F). These findings might be due to our experimental conditions, for example, the nominal Ca²⁺-free pipette solution, because the EC₅₀ values of riluzole for SKCa channels are reported to be affected by the intracellular Ca²⁺ concentration [45].

Given that the above-reported results indicate that TTX-R I_NaP plays a pivotal role in the various properties of the tested channels and the excitability of DiI-positive neurons, we examined whether peripheral sensitization with IMs affected the density of TTX-R I_NaP and neuronal excitability. A cocktail of IMs (5-HT, PGE₂, and bradykinin) with DiI was administered to the dura mater of living rats. After 7–10 days of this treatment, the DiI-positive neurons were used to perform an electrophysiological study (see Methods). In DiI-positive neurons isolated from Elvax-treated animals, the density of TTX-R I_NaP was correlated with neuronal size in both IM-treated and vehicle-treated groups, but the correlation coefficient was smaller in the IM-treated group (IM-treated group: r = 0.50, n = 30, p < 0.01; vehicle-treated group: r = 0.69, n = 42, p < 0.01; Fig. 7A). This was due to the increased density of TTX-R I_NaP recorded from small-sized DiI-positive neurons. Additionally, the mean density of TTX-R I_NaP recorded from small-sized DiI-positive neurons was significantly higher in the IM-treated group than that in the vehicle-treated group [3.8 ± 0.7 pA/pF (n = 17) vs. 7.9 ± 1.0 pA/pF (n = 15), respectively; p < 0.01; Fig. 7Ba]. In contrast, the density of TTX-R I_NaP recorded from medium-sized DiI-positive neurons did not differ significantly between the two groups [13.6 ± 1.2 pA/pF (n = 25) vs. 14.8 ± 1.4 pA/pF (n = 15), respectively; p = 0.50; Fig. 7Bb]. The mean density of TTX-R I_NaT did not differ between the IM-treated and vehicle-treated groups (data not shown). The density of TTX-R I_Ramp was correlated with neuronal size in both the IM-treated and vehicle-treated groups, but the correlation coefficient was smaller in the IM-treated group (IM-treated group: r = 0.55, n = 30, p < 0.01; vehicle-treated group: r = 0.59, n = 42, p < 0.01; Fig. 7C). The mean density of TTX-R I_Ramp recorded from small-sized DiI-positive neurons was significantly higher in the IM-treated group than that in the vehicle-treated group (4.5 ± 0.9 pA/pF (n = 17) vs. 9.7 ± 1.0 pA/pF (n = 15), respectively; p < 0.01; Fig. 7Da). However, the density of TTX-R I_Ramp recorded from medium-sized DiI-positive neurons did not differ statistically between the groups (p = 0.12; Fig. 7Db). Taken together, these results suggest that the noninactivating components of TTX-R I_Na were enhanced in small-sized DiI-positive neurons by peripheral sensitization due to IMs.

An increase in the density of TTX-R I_NaP is expected to alter the various properties of these channels in IM-treated small-sized DiI-positive neurons, as shown in the results reported above. Therefore, we also examined the voltage dependence, use-dependent inhibition, and inactivation and recovery kinetics of TTX-R Na⁺ channels in the IM-treated and vehicle-treated groups. We found that various properties of TTX-R Na⁺ channels were affected in small-sized but not medium-sized DiI-positive neurons (Fig. 7 and Supplementary Fig. S6). Although the V₅₀ values for activation did not differ between the groups, the V₅₀ values for fast inactivation were significantly shifted to the depolarizing range, where the mean V₅₀ values for fast inactivation obtained from small-sized DiI-positive neurons were -34.0 ± 1.3 mV (n = 17) and -39.3 ± 1.2 mV (n = 16) in IM-treated and vehicle-treated groups, respectively (p < 0.01; Fig. 7E). The mean V₅₀ values for slow inactivation obtained from small-sized DiI-positive neurons were also significantly higher in the IM-treated group compared to those in the vehicle-treated group [-44.7 ± 0.9 mV (n = 17) vs. -49.3 ± 1.5 mV (n = 15), respectively; p < 0.01; Fig. 7F]. The extent of use-dependent inhibition was reduced in the IM-treated group compared to that in the vehicle-treated group [P₂₀/P₁ ratio: 0.84 ± 0.02 for the IM-treated group (n = 13) vs. 0.74 ± 0.02 for the vehicle-treated group (n = 15); p < 0.01; Fig. 7G]. In addition, the inactivation observed from small-sized DiI-positive neurons developed significantly more slowly in the IM-treated group than in the vehicle-treated group. The τ_fast values were 418.2 ± 81.3 ms (n = 19) and 195.9 ± 23.5 ms (n = 16) in the IM-treated and vehicle-treated groups, respectively (p < 0.01; Fig. 7H, Ia); the τ_slow values were also larger in the IM-treated and vehicle-treated groups, respectively (p < 0.01, Fig. 7Ib). Furthermore, the extent of recovery from inactivation for small-sized DiI-positive neurons occurred significantly more quickly in the IM-treated group than in the vehicle-treated group; the τ_fast values were 6.9 ± 1.3 ms (n = 18) and 15.3 ± 3.2 ms (n = 15) in the IM-treated and vehicle-treated groups, respectively (p < 0.01; Fig. 7J, Ka). However, there was no difference in the τ_intermediate and τ_slow values between the IM-treated and vehicle-treated groups (Fig. 7Kb, Kc). In contrast, in medium-sized DiI-positive neurons, we detected no differences in voltage dependence, the onset of inactivation, and recovery from inactivation of TTX-R Na⁺ channels between the IM-treated and vehicle-treated groups (Supplementary Fig. S6).

Finally, we examined whether the excitability was changed in IM-treated DiI-positive neurons. The basal membrane properties of DiI-positive neurons in the vehicle-treated and IM-treated groups are summarized in Supplementary Table S2. While the firing patterns of small- or medium-sized DiI-positive neurons did not differ between the vehicle-treated and IM-treated groups, the rheobase currents measured in small-sized DiI-positive neurons were significantly lower in the IM-treated group than in the vehicle-treated group (Supplementary Table S2). In the presence of TTX (300 nM), action potentials were elicited by depolarizing current stimuli (1 T–4 T) in DiI-positive neurons of the vehicle-treated and IM-treated groups. In small-sized DiI-positive neurons, the number of action potentials was higher in the IM-treated group than in the vehicle-treated group (Fig. 8A). However, in medium-sized DiI-positive neurons, there was no difference in the number of action potentials observed between the vehicle-treated and IM-treated groups (Fig. 8B). We further examined the effects of riluzole on the excitability of IM-treated small-sized DiI-positive neurons. Riluzole (10 μM) greatly reduced the number of action potentials elicited by depolarizing current stimuli, in which it reduced the number of action potentials elicited by 4 T stimulation (11.8 ± 2.9 and 2.4 ± 0.4 under control and riluzole conditions, respectively, n = 8, p < 0.01, Fig. 8C). When single action potentials were elicited by brief depolarizing current stimuli (3 ms duration) in IM-treated small-sized DiI-positive neurons, riluzole (10 μM) had no effect on the amplitude of AHP (99.4 ± 1.0% of the control, n = 6, p = 0.48) and decay time constant of AHP (96.8 ± 3.6% of the control, n = 6, p = 0.45, Fig. 8D).

In the present study, the density of TTX-R I_NaP and TTX-R I_Ramp was correlated with the size of DiI-positive neurons. Both TTX-R I_NaP and TTX-R I_Ramp were sensitive to riluzole and A803467 (a specific Na_V1.8 blocker), indicating that these currents were mediated by the same channels. Such noninactivating currents might correspond to the I_NaP found in DRG neurons [21] as well as the central neurons in various brain regions [11, 46‐49]. The contamination of other TTX-R Na⁺ channel subtypes, such as Na_V1.9 or Na_V1.5, might be negligible because the V_{50,activation} and V_{50,inactivation} values for such subtypes have ranges that include relatively hyperpolarized potentials [18, 50]. In addition, since the voltage-fast inactivation relationships of TTX-R I_NaT and TTX-R I_NaP were largely the same, TTX-R I_NaP is mediated by Na_V1.8 rather than other subtypes. Consistent with these results, we also found that the voltage dependences of TTX-R Na⁺ channels, such as the activation and instantaneous steady-state fast inactivation relationships, did not differ among DiI-positive neurons and that relationships did not exist between the voltage dependences and neuronal size or the density of TTX-R I_NaP. However, there were marked differences in the fast component (i.e., τ_fast) of slow inactivation and in recovery kinetics of small- and medium-sized DiI-positive neurons. The τ_fast values for the onset of slow inactivation were significantly lower in small-sized DiI-positive neurons than in medium-sized neurons, and, importantly, these values were correlated with the density of TTX-R I_NaP and, to a lesser extent, neuronal size. Similarly, the τ_fast values for recovery from inactivation were significantly higher in small-sized DiI-positive neurons than those in medium-sized neurons, and these values were also correlated with the density of TTX-R I_NaP. Thus, TTX-R I_NaP are apparently involved in channel properties such as slow inactivation and recovery kinetics.

In the current study, changes to the holding potential and recovery duration did not affect the τ_fast values in both small- and medium-sized DiI-positive neurons. Because large differences in τ_fast values existed between small- and medium-sized DiI-positive neurons, the holding potential and/or recovery duration might not be responsible for the fundamental difference in the fast component of inactivation kinetics. Observed changes in A_fast values suggest that the holding potential and recovery duration may affect the extent rather than the rate of slow inactivation. We also found that the τ_fast values for recovery kinetics were greatly decreased or increased by shortening or lengthening the duration of the inactivating prepulse, respectively, in small- and medium-sized DiI-positive neurons, suggesting that the extent of slow inactivation during depolarizing prepulses might determine the recovery kinetics of TTX-R Na⁺ channels in these neurons. Therefore, differences in the slow inactivation of TTX-R Na⁺ channels are likely to result in marked differences in recovery kinetics and use-dependent inhibition, as shown by other researchers [9, 10]. Notably, the extent of use-dependent inhibition and kinetics for slow inactivation and recovery from inactivation of TTX-R Na⁺ channels were more strongly correlated with the density of TTX-R I_NaP than with cell capacitance, suggesting that a relationship potentially exists between TTX-R I_NaP and channel functions.

Growing evidence supports the important role played by I_NaP in the repetitive firing of action potentials in response to sustained membrane depolarization in central neurons [11, 51‐56]. In addition, the amplitude of I_NaP is known to be related to the firing patterns of spinal neurons [57]. Human Na_V1.8 produces larger persistent currents and ramp currents than those produced by rat Na_V1.8, and these properties of Na_V1.8 currents are related to the increased firing frequency in neurons [21]. However, the functional role of TTX-R I_NaP in nociceptive neurons remains largely unknown. In our study, the firing patterns and number of action potentials in response to depolarizing current stimuli differed considerably between the two tested types of DiI-positive neurons. Given that firing patterns were not changed even in the presence of TTX and that riluzole greatly reduced the extent of the repetitive generation of action potentials in small- and medium-sized DiI-positive neurons, riluzole-sensitive TTX-R I_NaP might play a major role in the repetitive generation of action potentials in C-type dural afferent neurons.

Several factors may be responsible for the differential density of TTX-R I_NaP and inactivation kinetics in C-type dural afferent neurons. For example, the incorporation of accessory β subunits into various subtypes of Na⁺ channels is known to affect I_NaP and inactivation kinetics [58]. In a previous study using Scn1b^−/− mice, β1 subunits contributed to larger TTX-R I_NaP and faster recovery from inactivation in small DRG neurons [59]. However, since a marked difference in the inactivation kinetics between Scn1b^−/− and Scn1b^+/+ mice was not detected [59], β1 subunits alone are unlikely to be responsible for the different TTX-R Na⁺ channel properties among dural afferent neurons. Alternatively, splice variants for Na_V1.8 [60, 61] might explain the marked differences in the TTX-R Na⁺ channel properties of small- and medium-sized DiI-positive neurons. However, the obvious impact of splicing events on the function of Na_V1.8 has yet to be found [61]. Thus, further research is required to reveal which factors are involved in the differential properties of TTX-R Na⁺ channels in C-type dural afferent neurons. Nevertheless, previous studies have demonstrated the differential slow inactivation and use-dependent inhibition of Na_V1.8 in two subpopulations of small-sized DRG neurons, i.e., isolectin B₄-positive nonpeptidergic and IB₄-negative peptidergic neurons [10]. Furthermore, the protein calmodulin, which is reportedly involved in the TTX-R Na⁺ channel differs between these two subpopulations [62]. However, in the present study, most small- and medium-sized DiI-positive neurons were likely to be peptidergic neurons because ~ 80% of these neurons express CGRP [23], which is regarded as a peptidergic neuron marker [63]. It would be of interest to examine whether calmodulin is also involved in the differential properties of TTX-R Na⁺ channels in C-type dural afferent neurons.

The increased neuronal excitability of nociceptive neurons is closely related to inflammatory hyperalgesia, i.e., IM treatment increases the number of action potentials elicited by depolarizing current stimuli in nociceptive neurons [23, 25]. In this regard, Na_V1.8 expressed in nociceptive neurons are reportedly subject to inflammatory sensitization because these channels are positively modulated by IMs [64, 65]. Furthermore, CGRP-induced neurogenic inflammation and the subsequent sensitization of dural afferent neurons play a pivotal role in migraine pathology [66, 67]. In the present study, we found that the density of TTX-R I_NaP was significantly larger in small-sized DiI-positive neurons treated with IM than those treated with a vehicle. This increase in TTX-R I_NaP density could lead to changes in channel properties, including the inactivation and recovery kinetics, which might eventually affect the firing properties and/or frequency of action potentials in relation to depolarizing stimuli. Interestingly, mutations in Na_V1.1, which are associated with familial hemiplegic migraine type 3, cause an increase in the density of I_NaP, a defect in the inactivation process, and repetitive generation of action potentials [68]. Although the mechanisms that underlie the IM-induced increase in TTX-R I_NaP remain to be elucidated, our study suggests that TTX-R I_NaP are involved in peripheral sensitization mediated by neurogenic inflammation around the dura mater. However, in medium-sized DiI-positive neurons, the density of TTX-R I_NaP and the properties of TTX-R Na⁺ channels did not differ between the IM-treated and control groups. Therefore, further studies are required to determine the role of medium-sized DiI-positive neurons in the inflammatory sensitization of dural afferents.

Voltage-gated Na⁺ channels may be related to migraine pathology, given that most of the effective drugs used for migraine prophylaxis, such as β-blockers (e.g., propranolol), tricyclic antidepressants (e.g., amitriptyline), Ca²⁺ channel blockers (e.g., flunarizine), and Na⁺ channel blockers (e.g., valproic acid) [69, 70], are known to inhibit voltage-gated Na⁺ channels [71‐74]. Although there is no direct evidence that TTX-R Na⁺ channels and TTX-R I_NaP are involved in migraine pathology, our previous studies have also shown that either amitriptyline or propranolol preferentially inhibit TTX-R I_NaP and decrease the excitability of dural afferent neurons [75, 76]. It would be interesting to investigate whether CGRP-mediated neurogenic inflammation affects the properties of TTX-R Na⁺ channels and TTX-R I_NaP and whether drugs used to treat and prevent migraine attacks can inhibit TTX-R I_NaP in dural afferent neurons.

In the present study, we shed light on the role of Na_V1.8-mediated I_NaP in the excitability of dural afferent neurons. However, nociceptive neurons also express another TTX-R Na⁺ channel subtype, Na_V1.9 [18.19]. Na_V1.9 has been implicated in inflammatory hyperalgesia induced by IMs [77‐79], and Na_V1.9-mediated currents are potentiated by IMs [20]. Interestingly, a recent study reported that with the abnormal activation of Na_V1.9, the TTX-R Na⁺ channels subtypes expressed in nociceptive neurons by nitric oxide are responsible for medication-overuse headaches induced by triptan migraine medicine [80]. Since Na_V1.9-mediated currents, including slow voltage ramp-induced currents, were recorded from most small- and medium-sized DiI-positive neurons, it would be of great interest to examine whether Na_V1.9-mediated I_Ramp is also involved in the IM-mediated increase in the excitability of small-sized DiI-positive neurons.