Background
Substance P (SP), a member of tachykinin family, is a well-known pain-related neuropeptide in the spinal cord. It is released by unmyelinated primary afferent fiber terminals of small-diameter dorsal root ganglion (DRG) neurons and participates in the spinal transmission of nociceptive signals [
1‐
3]. It is well documented that the SP receptor neurokinin-1 (NK-1) is densely distributed in the superficial dorsal horn and involved in the development of chronic pain and central sensitization after intense noxious stimulation and tissue/nerve injury [
4‐
7].
In addition to the expression of the NK-1 on the postsynaptic neurons of superficial spinal dorsal horn, increasing evidence strongly suggested the presynaptic expression of NK-1 in DRG neurons. The immunohistochemical evidence revealed that the NK-1 was expressed by the unmyelinated axons of the glabrous skin [
8], and the DRG neuron soma in rats [
9]. By means of intracellular and whole-cell patch clamp recordings, SP was shown to be able to induce the depolarization of DRG or trigeminal ganglion neurons in the different species [
10‐
13] and potentiated the TRPV1 currents [
9]. However, the function of DRG-expressed NK-1 receptor needs to be further understood.
The Na
v1.8, which is a TTX-resistant sodium channel and mainly expressed in small-diameter DRG neurons [
14,
15], is a major contributor to the upstroke of action potential in these neurons [
16]. In the Na
v1.8-null mice or Na
v1.8 knockdown mice by antisense oligodeoxynucleotides, both the physiological and pathological pain was alleviated [
17‐
20]. Accumulative evidence showed that the Na
v1.8 current was regulated by various inflammatory mediators, such as prostaglandin E
2 (PGE
2), serotonin, NGF etc. through a PKA or PKC signaling pathway [
21‐
24].
In the present study, we investigated the effects of the NK-1 agonist on dynamics of Nav1.8 currents in isolated small-diameter DRG neurons using whole-cell patch clamp recording. Also, the role of PKC signal pathway in the cross-talk between NK-1 and Nav1.8 was examined.
Discussion
The role of substance P (SP) and its NK-1 receptor in pain processing was widely investigated in the spinal cord. However, most of the prevailing studies focused on the postsynaptic NK-1 receptors in the spinal superficial dorsal horn neurons. Whether NK-1 receptors are also expressed presynaptically in primary sensory neurons is still obscure. A growing body of evidence showed that SP could activate DRG neurons through NK-1 receptor in primary sensory neurons [
10‐
13,
34,
35], in despite of a contradictory report [
36]. Our recent study provided new evidence for the expression of NK-1 receptor protein and interaction with TRPV1, a crucial pain signal molecule, in DRG neurons [
9].
In addition to TRPV1, another important ion channel, TTX-resistant sodium channel, is also primarily expressed in nociceptors. Between the two distinct TTX-resistant sodium channel isoforms Na
v1.8 and Na
v1.9, Na
v1.8 likely mediates the majority of the TTX-resistant currents and plays an important role in pain processing. Na
v1.8-null mice displayed a pronounced increase in threshold to noxious mechanical stimuli and a slight decrease in nociceptive thermoreception as well as delayed development of inflammatory hyperalgesia [
17]. Likewise, knocking-down of Na
v1.8 mRNA with antisense oligodeoxynucleotides was effective in alleviating both the inflammatory and neuropathic pain [
18‐
20,
37,
38]. Also, muO-conotoxin MrVIB, a selective blocker of Na
v1.8, reduced allodynia and hyperalgesia in neuropathic and chronic inflammatory pain models [
39,
40]. The present study for the first time revealed that NK-1 activation potentiated Na
v1.8 currents and shifted both the activation and steady-state inactivation curves of this channel in a hyperpolarizing direction. This change in voltage sensitivity of Na
v1.8 may decrease the activate threshold and increase the likelihood of action potential firing, and then probably cause a hyperexcitability of the neurons. As shown in Figure
6, the enhancement of excitability was observed in our experiments. Although the involvement of Sar-SP-induced modulation on other ion channels still need to be further investigated, it is assumed that modulation on Na
v1.8 at least partly contributes to this enhancement of excitability. Similar results were also obtained from studies on modulation of Na
v1.8 by another peripheral pain-related neuropeptide calcitonin gene related peptide (CGRP) [
41] and proinflammatory factors such as 5-hydroxytryptamine and prostaglandin E
2 [
22,
42,
43]. It is conceivable that the modulation of Na
v1.8 by NK-1 activation may contribute to peripheral sensitization of pain pathway.
NK-1 receptor is a G-protein coupled receptor [
28]. The activation of NK-1 receptors generates various second messengers, which, in turn, trigger a wide range of effector mechanisms underlying regulating cellular excitability and functions [
44‐
47]. In agreement with our previous finding that the modulation of TRPV1 by NK-1 receptor was mediated by activation of PLC and downstream PKC pathway [
9], the present results also proved the involvement of PKC in the interaction between NK-1 and Na
v1.8. As shown in Figure
3 and Figure
4, PKC inhibitor BIM completely blocked Sar-SP-induced potentiation of Na
v1.8 currents, whereas PKC activator PMA could mimic the effects of Sar-SP on Na
v1.8 currents. These results suggest that NK-1 modulates Na
v1.8 in a PKC-dependent pathway. There are many other papers confirmed the enhancement of Na
v1.8 by PKC pathway [
24,
48,
49]. However, the inconsistent results have been reported. Gold et al. reported that PKC activation also caused an increase in the amplitude of the TTX-resistant current in rat DRG neurons. But this increase was not associated with a shift in the activation curve [
23]. Vijayaragavan and colleagues reported that in
Xenopus oocytes expression system, PKC activator PMA caused a decrease of Na
v1.8 current and a right shift of the activation curve [
29]. The reason for the difference is still unclear.
Furthermore, we observed that PKCε inhibitor εV1-2 completely blocked Sar-SP-induced potentiation of Na
v1.8 currents, suggesting that PKCε was the main mediator of NK-1 potentiation, in consistence with the modulation of TRPV1 by NK-1 [
9].
In addition to the PKC pathway, several reports showed that PKA was also involved in the increase of TTX-resistant currents by proinflammatory agents (5-HT, PGE
2) [
22,
23,
42]. However, PKA inhibitor H89 failed to prevent Sar-SP-induced potentiation of Na
v1.8 in the present study. These suggested the diverse mechanisms underlying modulation of Na
v1.8 by the different proinflammatory agents. Therefore, the modulatory action of NK-1 may be predominately mediated by PKC, particularly by PKCε, but not PKA.
It is well documented an increase in expression of Na
v1.8 in DRG neurons in several inflammatory pain models [
50‐
53]. Our previous results have revealed that both the NK-1 expression and phosphorylation of PKCε are up-regulated in DRG after CFA-induced inflammation [
9,
33]. Therefore, we assume that the modulation of Na
v1.8 by NK-1 via PKCε is likely to be stronger after peripheral inflammation. In support of this view, the present study showed that not only the effect of NK-1 activation on Na
v1.8 currents was significantly potentiated, but also the rate of Sar-SP-responsive neurons following CFA treatment. It is conceivable that the modulation of Na
v1.8 by NK-1 may amplify peripheral nociceptive inputs and in turn strengthen activation of the pain-sensitive neurons in the spinal cord, contributing to inflammatory pain.
Methods
Animals
Male adult (100–150 g) Sprague-Dawley rats (obtained from the Experimental Animal Center, Shanghai Medical College of Fudan University, China) were used in our experiments. Rats were on a 12 h light/dark cycle with a room temperature of 22 ± 1°C and received food and water ad libitum. All experimental procedures were approved by the Shanghai Animal Care and Use Committee and followed the policies issued by the International Association for the Study of Pain on the use of laboratory animals. All efforts were made to minimize animal suffering and reduce the numbers of animals used.
Cell preparation
Culture of DRG neurons was established as described previously [
33]. Briefly, DRGs from L
4-L
6 lumbar segments were dissected and incubated at 36.8°C for 25 min in DMEM containing 3 mg/ml collagenase (type IA, Sigma, St. Louis, MO) and, 1 mg/ml trypsin (type I, Sigma). The ganglias were then gently triturated using fine fired-polished Pasteur pipettes. The dissociated DRG neurons were plated onto coverslips (10 mm diameter) in the 3.5 cm culture dishes and incubated with Standard external solution containing (in mM) 150 NaCl, 5 KCl, 2 CaCl
2, 1 MgCl
2, 10 HEPES, and 10 glucose, adjusted to pH 7.4 with NaOH.
Patch-clamp recordings
Whole-cell voltage-clamp and current-clamp recordings of DRG neurons were performed at room temperature (20–22°C) with an EPC-9 amplifier (HEKA Elektronik, Lambrecht/Pfalz, Germany). Stimulation protocols and data acquisition were controlled by the software Pulse and Pulsefit 8.5 (HEKA Elektronik). Neurons were prepared as above, and all recordings were performed within 2–8 h after plating. All of the recordings were made from small-diameter (15–25 μm) DRG neurons. After gigaohm seal formation and membrane disruption, the whole cell capacitance was cancelled and series resistance was compensated (> 80%). Microelectrodes were fabricated from 1.5 mm out diameter borosilicate capillary glass (Sutter Instruments, Novato, CA) by using a P-97 puller (Sutter Instruments, Novato, CA), and had a resistance of 3–5 MΩ. Electrodes were filled with (in mM): 140 CsF, 1 MgCl2, 1 EGTA, 2.5 Na2ATP, 10 HEPES, pH was adjusted to 7.2 with CsOH. In recording of Nav1.8 currents, the external solution contained (in mM): 32 NaCl, 20 TEA-Cl, 105 choline-Cl, 1 MgCl2, 1 CaCl2, 0.1 CdCl2, 10 HEPES, 0.0005 TTX and 10 glucose, adjusted to pH 7.4 with NaOH. The TEA-Cl, CdCl2, TTX was used to inhibit endogenous K+, Ca2+, and TTX-sensitive sodium currents, respectively. In current-clamp recordings, the electrode solution was changed to: 140 KCl, 1 MgCl2, 0.5 CaCl2, 5 EGTA, 10 HEPES, 2.5 Na2ATP, pH was adjusted to 7.2 with KOH. The external solution was changed to: 150 NaCl, 5 KCl, 2.5 CaCl2, 1 MgCl2, 10 HEPES, pH was adjusted to 7.4 with NaOH.
Drugs
All the drugs were purchased from Sigma (St. Louis, MO, USA), except that the PKCε inhibitor εV1-2 and its negative control were from Biomol (Plymouth Meeting, PA). All the drugs are prepared on the day of the experiment from stocks kept at -20°C at a concentration at least 1000-fold the working concentration. [Sar9, Met(O2)11]-substance P (Sar-SP) and PMA were applied close to the cells through a ALA-VM8 perfusion system (ALA Scientific Instruments, Westbury, NY). Inhibitors were applied (where appropriate) to the chamber for 30 min before the perfusion of Sar-SP and PMA and existed during the whole recording course.
Data analysis
Peak sodium current values were converted to conductance values using the equation: G = I/(V
m - E
Na), where G is the conductance, I is the peak current amplitude, V
m is the membrane potential, and E
Na is the equilibrium potential value for Na+. The Boltzmann equation used to describe the voltage dependence of activation was of the form: G/G
max
= 1/(1 + exp [(V
1/2 - V
m
)/k]), where G
max is the peak conductance, V
1/2 is the potential at half maximal activation, and k is the slope factor. Voltage dependence of steady-state inactivation was described by the Boltzmann function: I/I
max
= 1/(1 + exp [(V-V
1/2)/k]), where I
max is the maximal peak current, V is the prepulse membrane potential, V
1/2 is the potential at half maximal activation, and k is the slope factor.
Data are expressed as means ± standard error of the mean (SEM). Statistical analysis was performed using SigmaStat software (Systat Software, Chicago, IL). Student's t-test or Mann-Whitney analysis was used to assess differences between means from two groups. One-way ANOVA or Kruskal-Wallis one-way ANOVA was used to assess difference among more groups. P < 0.05 was considered significant. Curves were plotted and fitted using Origin software (OriginLab Corporation, Northampton, USA).
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
CLC and HZ performed the patch clamp recordings in DRG neurons. YQZ was partially involved in experimental design and guiding. ZQZ is the corresponding author.