This article is mentioned in:

Introduction

Neuropathic pain is an unmet medical concern, affecting individuals globally. The condition has been causally correlated with functional alterations in the sodium channels of nociceptors (1). Sodium channels are integral membrane glycoproteins, which are responsible for the generation and conduction of action potentials in excitable cells (2). Previous studies have revealed that sodium channel blockers, including local anesthetics, tricyclics and certain anti-convulsants, are able to attenuate pain in patients with neural injury (3,4). The voltage-gated sodium channel isoforms, Nav1.8 and Nav1.9, encoding for slowly-gating tetrodotoxin-resistant (TTX-R) sodium currents, are particularly notable with respect to sensory nerve pathophysiology. They are sensory neuron-specific with Nav1.8 expressed in thinly unmyelinated (C-fibers) and 10% myelinated (A-fibers) axons. By contrast, the expression of Nav1.9 is restricted to small C-fiber dorsal root ganglion (DRG) cells (5). Differential expression of these sodium channels is coupled with isoform-specific contributions in neuronal excitability and the transmission of sensory information (6). Nav1.8 produces the majority of the depolarizing inward current during an action potential (7), while Nav1.9 has been proposed to contribute to maintaining the resting potential (8). The absence of these channels in the central nervous system implicates them as a suitable target for therapeutic intervention in pain management with few side effects (2).

Peripheral nerve injury, for example axotomy or nerve transection, causes a downregulation of Nav1.8 expression and a decrease in the electrical current attributed to this channel in injured neurons (3,9,10). Additionally, specific knockdown of Nav1.8 with antisense oligodeoxynucleotides may effectively reverse neuropathic pain (11). However, it is not intuitively clear how this contributes to the neuropathic pain phenotype associated with these models. Previous studies have observed that upregulation of the expression of Nav1.8 in spared neurons following nerve injury, with the exception of chronic constriction injury (CCI) or spared nerve injury models, may provide a reasonable explanation for the contribution of this channel to the pain phenotype (10,12–14). In addition, there are specific discrepancies in the evidence among studies with regards to the involvement of Nav1.9 in certain neuropathic pain conditions (2,15–17). The expression of Nav1.9 is downregulated in injured DRG neurons (15) and is upregulated in uninjured neurons following peripheral nerve injury (16), while abnormal behavior in the Nav1.9 knockout mouse remains unchanged in neuropathic pain models (2). To resolve this limitation, a CCI model was established in the present study to evaluate whether Nav1.8 sodium channels mediate neuropathic pain through a compensatory redistribution to contralateral uninjured DRG neurons and to elucidate the exact role of Nav1.9 in neuropathic pain.

Materials and methods

Experimental animals

A total of 72 adult male Sprague-Dawley rats weighing 150–180 g and aged 6–8 weeks, purchased from the Experimental Animal Centre of Shenyang Pharmaceutical University (Shenyang, China), were used in all experiments. Rats were maintained under controlled environmental conditions at 23±2°C with a 12-h light/dark cycle and ad libitum access to food and water. All procedures were performed in accordance with the guidelines of the International Association for the Study of Pain (18). The study was approved by the ethics committee of Shenyang Pharmaceutical University.

CCI model

The CCI model was induced as previously described (19,20). Briefly, under anesthesia with 3.5% chloral hydrate (10 ml/kg; Hebei Gaobeidian Chunguang Chemical Reagent Company, Gaobeidian, China) administered intraperitoneally (i.p.), the right sciatic nerve was exposed and loosely ligated with 4–0 chromic catgut (Shanghai Pudong Medical Supplies Co., Ltd., Shanghai, China) in four regions separated by ~1 mm, then the incision was closed with sutures. For the sham group, the right sciatic nerve was exposed without ligation. Pain-associated behavioral assessments were performed at the time-points of: −1 (prior to CCI surgery), 1, 3, 5, 7, 10, 14 and 21 days after CCI surgery.

Behavioral assessment

The abnormal posture of each animal was evaluated using a subjective pain-associated behavioral grade as described previously (21) by an investigator who was blinded to the experimental conditions. Briefly, grades were determined as: 0, normal; 1, coiling of the toes; 2, valgus deformity of the paw; 3, partially weight bearing; 4, non-weight bearing and 5, avoidance of any contact with the hind paw.

Paw withdrawal response to thermal stimuli

The paw withdrawal response was measured using the Plantar Test meter (IITC Life Science Inc., Woodland Hills, CA, USA). Rats were placed individually in a clear, transparent box (17×11.5×14 cm) and allowed to acclimate for 15 min prior to the assessment. An infrared beam of a radiant heat source was applied to irradiate the plantar surface of the hind paw through the glass plate until the rat withdrew or contracted its paw. The withdrawal thresholds were measured in each hind paw and the ipsilateral hind paw was assessed at 15 min intervals, while the contralateral hind paw was assessed at 5 min intervals. A 20 sec limit was imposed to prevent tissue damage. Each rat was assessed five times and the mean value was expressed as the thermal withdrawal latency (22).

The paw withdrawal response to mechanical stimuli was measured using the Electronic von Frey Anesthesiometer (IITC Life Science Inc.). Rats were placed individually into wire mesh-bottom boxes (20×14×16 cm) and allowed to acclimate for 30 min prior to assessment. The probe was placed beneath the plantar surface of the hind paw and the force was increased until the rat was observed to vellicate its paw. The maximum force was recorded at the time of paw withdrawal. Withdrawal thresholds were measured in each hind paw and the ipsilateral hind paw was assessed at 30 sec intervals, while the contralateral hind paw was assessed at 15 sec intervals. Each rat was assessed five times and the average value was expressed as the mechanical withdrawal threshold (23).

Immunofluorescence and hematoxylin and eosin (H&E) staining

L4-6 DRGs, quickly dissected from recently sacrificed animals, were fixed with 10% formalin overnight at 4°C. Following embedding the tissue in paraffin, a series of 5-µm sections were cut for immunofluorescence and H&E staining. Paraffin sections were treated with dimethylbenzene solution I for 15 min, dimethylbenzene solution II for 15 min, dimethylbenzene:pure ethanol (1:1) solution for 2 min, 100% ethanol I for 5 min, 100% ethanol II for 5 min, 95% ethanol solution for 3 min, 90% ethanol solution for 1 min, 85% ethanol solution for 1 min, 75% ethanol solution for 1 min, 50% ethanol solution for 1 min, running water for 2 min, haematoxylin solution for 2 min, running water for 1 min, 1% hydrochloric acid ethanol solution for 20 sec, running water for 5 min, Eosin solution for 30 sec, running water for 30 sec, 75% ethanol solution for 30 sec, 85% ethanol solution for 20 sec, 95% ethanol solution I for 1 min, 95% ethanol solution II for 1 min, 100% ethanol for 2 min, 100% ethanol II for 2 min, dimethylbenzene solution I for 2 min, dimethylbenzene solution II for 2 min then dimethylbenzene solution III for 2 min. The primary antibodies polyclonal rabbit Nav1.8 (ASC-016; 1:200; Alomone Laboratories Ltd., Jerusalem, Israel), monoclonal mouse neurofilament (NF)200 (N0142; 1:200; Sigma-Aldrich, St. Louis, MO, USA) and polyclonal rabbit Nav1.9 (ASC-017; 1:200; Alomone Laboratories Ltd.) were administered for immunofluorescence staining and used to incubate the sections overnight at 4°C. Following three washes with phosphate-buffered saline containing Tween-20, the sections were incubated with anti-mouse IgG fluorescein isothiocyanate-conjugated antibody produced in goat (F0257) and anti-rabbit IgG (whole molecule)-Cy3 antibody produced in sheep (C2306) (1:100; Sigma-Aldrich) for 1 h at 37°C. Images were captured under an inverted fluorescence microscope (Olympus BX40; Olympus, Tokyo, Japan) and imported into Image pro plus 6.0 software (Media Cybernetics, Silver Spring, MD, USA) for further analysis. H&E staining was performed according to the manufacturer's instructions (Sigma-Aldrich). Nuclear material within the nucleus was stained a deep purple/blue, while the cytoplasmic material, including connective tissue and collagen appeared orange/pink.

Reverse transcription-quantitative polymerase chain reaction (RT-qPCR)

At 3, 7, 14 and 21 days after CCI, rats were sacrificed via anesthesia with 3.5% chloral hydrate (10 ml/kg, i.p.). The ipsilateral and contralateral L4–6 DRGs were rapidly removed and placed into Eppendorf tubes. Total RNA was extracted using TRIzol reagent (Invitrogen Life Technologies, Carlsbad, CA, USA) and the cDNA was reverse transcribed using the PrimeScript® RT reagent kit (Takara Bio Inc., Otsu, Japan) according to the manufacturer's instructions. PCR was performed in a 25-µl reaction mixture containing 2 µl templates, 12.5 µl SYBR® Premix Ex Taq™ (2X), 0.5 µl ROX reference dye II (50X) and 0.4 µM primer for each gene. The thermal cycling conditions comprised 30 sec polymerase activation at 95°C, 40 cycles of 15 sec denaturation at 95°C and 1 min at 60°C for annealing and extension. A dissociation curve was used to determine the amplification specificity. The primer sequences (24) were as follows: Nav1.8 (GenBank accession number, U53833) forward, 5′-GACTCCCGGACAAATCAGAA-3′ and reverse, 5′-AGCAGCGACCTCATCTTCAT-3′; Nav1.9 (GenBank accession number, AF059030) forward, 5′-TCTCCACCCCTACCTCACTG-3′ and reverse, 5′-CGTTCAGCCAAAAACACAGA-3′; and GAPDH forward, 5′-TGCCAAGTATGATGACATCAAGAAG-3′ and reverse, 5′-AGCCCAGGATGCCCTTTAGT-3′. The threshold cycle values of Nav1.8 and Nav1.9 mRNA were measured and normalized to GAPDH, and then expressed as a relative ratio. The M×3000P qPCR system was used (Agilent Technologies, Inc., Santa Clara, CA, USA).

Western blot analysis

Bilateral L4–6 DRGs were dissected and total proteins were extracted via homogenization in ice-cold lysate buffer (Thermo Fisher Scientific, Waltham, MA, USA). Samples (30–50 µg) were separated on 8% SDS-PAGE separation gels (Amresco, Boise ID, USA) and subsequently transferred onto polyvinylidene difluoride membranes (Merck Millipore, Boston, MA, USA). The membranes were blocked in 5% skimmed milk solution at room temperature for 2 h and then rabbit polyclonal Nav1.8 (1:500; Alomone Laboratories Ltd.), Nav1.9 (1:500; Alomone Laboratories Ltd.) and mouse monoclonal β-actin (1:500; Santa Cruz Biotechnology, Inc., Dallas, TX, USA) primary antibodies were used to incubate the samples overnight at 4°C. The target bands were detected with secondary horseradish peroxidase (HRP)-labeled goat anti-rabbit (1:10,000; ZB-2301) or HRP-labeled goat anti-mouse IgG (1:5,000; ZB2305) (Zhongshan Golden Bridge Biotechnology Co., Ltd., Beijing, China) antibodies for 1 h at room temperature. The band intensity of Nav1.8 and Nav1.9 was normalized to that of β-actin and expressed as a relative ratio.

Patch clamp recording

Rat DRG neurons were acutely dissociated as previously described (25). The samples were superfused at a rate of 3 ml/min and all patch clamp recordings were performed using an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, CA, USA), filtered at 1 kHz and digitally sampled at 10 kHz at room temperature (23±2°C). Clampfit 10.0 software (Molecular Devices) was used for data acquisition and analysis.

The bath solution for the Nav1.8 currents contained (in mM): 140 NaCl, 1 MgCl2, 3 CaCl2, 5 KCl (Tianjin Bodi Chemical Co. Ltd., Tianjin, China), 10 tetraethylammonium (TEA)-Cl, 1 4-aminopyridine, 0.2 CdCl2, 10 4-(2-hydroxyethyl)-1-piper-azineethanesulfonic acid (HEPES), 10 glucose (Tianjin Bodi Chemical Co. Ltd.), 0.001 TTX (Hebei Fisheries Research Institute, Qinhuangdao, China), pH 7.3 with NaOH (Tianjin Bodi Chemical Co. Ltd.). The bath solution for the Nav1.9 currents contained 30 NaCl, 20 TEA-Cl, 90 choline chloride, 3 KCl, 1 CaCl2, 1 MgCl2, 10 HEPES, 10 glucose, 0.1 CdCl2, 0.001 TTX, pH 7.3 with Tris base (all purchased from Sigma-Aldrich unless specified). The pipette solution used for recording the DRG neuron Nav1.8 and Nav1.9 currents contained (in mM): 140 CsCl, 10 TEA-Cl, 10 ethylene glycol tetraacetic acid (EGTA; Amresco), 10 HEPES, pH 7.2 with CsOH and 135 CsF, 10 NaCl, 10 HEPES, 5 EGTA, 2 adenosinetriphosphate bisodium, pH 7.2 with CsOH, respectively. The pipettes, fabricated with a P-97 puller (Sutter Instruments, Novato, CA, USA), had resistances of 3–5 MΩ when filled with pipette solution.

Statistical analysis

Data were analysed using SPSS 16.0 (SPSS, Inc., Chicago, IL, USA) with a one-way analysis of variance. All measurements are expressed as the mean ± standard error. P<0.05 was considered to indicate a statistically significant difference.

Results

CCI-induced neuropathic pain evokes spontaneous pain, mechanical allodynia and heat hyperalgesia

The CCI model induced the typical features of neuropathic pain, which were assessed via behavioral grade, the Plantar Test meter and the Electronic von Frey Anesthesiometer. Rats with nerve injury (CCI) exhibited curling, eversion, partial weight bearing or non-weight bearing on the right injured side, while they exhibited normal behavior on the left uninjured paw. The score for the right paw increased significantly from the first day after surgery (data not shown).

The sensitivity to heat and mechanical stimulus revealed certain discrepancies among the experimental groups. In the CCI model group, the thermal withdrawal latency and mechanical withdrawal threshold of the right injured paw decreased markedly from day 1 after injury, with the maximal level of hypersensitivity at 7 days after surgery, while observable changes were not observed in the sham surgery group (Fig. 1A and C). By contrast, no differences were observed in the contralateral paw withdrawal in all groups (Fig. 1B and D).

Figure 1 CCI-induced neuropathic pain evokes heat hyperalgesia and mechanical allodynia. (A and C) Significant decreases in the ipsilateral mean paw withdrawal latency and mean paw withdrawal threshold were observed from day 1 and persisted for 21 days. (B and D) Contralateral paw withdrawal. Values are expressed as the mean ± standard error of the mean. *P<0.05; ***P<0.001 vs. sham group. CCI, chronic constriction injury.

CCI-induced neuropathic pain impairs DRG histological morphology

In peripheral nerve injury, the majority of neuronal cell death is observed in peripheral nerve lesions, which are considered to be a major factor in the poor clinical outcome following the injury (26). With H&E staining, ipsilateral and contralateral DRGs to the injury (Fig. 2) were analyzed histologically. In each contralateral DRG group (Fig. 2G–L), the neurons exhibited a normal cellular appearance, including a large, round nucleus, clear cell boundaries and even chromatin distribution. However, neurons on the ipsilateral side to the injury exhibited a range of morphologies at different time-points (Fig. 2C–F), although all observed cells had abnormal nuclei exhibiting pyknosis, membrane irregularities or even vacuolation, with the maximal magnitude of neuronal death at 21 days after surgery.

Figure 2 Micrographs of H&E staining in representative L4–6 DRG slices. Representative sections stained with H&E revealed a gradual injury (arrow) in ipsilateral DRG cells at the point of (C) 3, (D) 7, (E) 14 and (F) 21 days after CCI compared with the sham group (B). Normal cellular morphology is shown in (A) the control and (B) the sham group ipsilateral and (G–L) in the other groups' contralateral L4–6 DRG of rats. Scale bar, 50 µm. H&E, hematoxylin and eosin; DRG, dorsal root ganglion; CCI, chronic constriction injury.

Expression profiles of Nav1.8 and Nav1.9 in DRG neurons under CCI-induced neuropathic pain

The neurofilament NF200 is preferentially expressed in large DRG neurons and is a useful marker for this population (7). When co-incubated with antibodies against Nav1.8 and Nav1.9 in an immunofluorescence assay, it was observed that Nav1.8 and Nav1.9 subunits were primarily located in the small and medium diameter DRG neurons (Fig. 3A).

Figure 3 Changes in the expression and distribution of Nav1.8 and Nav1.9 following CCI. (A) DRG sections were labeled with anti-NF200, anti-Nav1.8 and anti-Nav1.9 antibodies. (B) Nav1.8 mRNA levels were significantly decreased in ipsilateral DRGs at 7, 14 and 21 days, while Nav1.9 expression was markedly lower at 14 and 21 days after CCI. (C) Relative expression of Nav1.8 and Nav1.9 mRNA in contralateral DRGs. (D) Representative images of Nav1.8 and Nav1.9 protein and (E and F) relative ratio of Nav1.8 protein exhibited a significant decrease at 7–21 days after CCI in ipsilateral DRGs. Values are expressed as the mean ± standard error of the mean. **P<0.01; ***P<0.001, vs. sham group. DRG, dorsal root ganglion; CCI, chronic constriction injury; NF200, neurofilament 200.

RT-qPCR and western blot analyses were performed at 3, 7, 14 and 21 days after the CCI surgery instead of at an earlier point, in order to avoid potential error due to the effects of post-surgical pain. CCI induced a marked downregulation of the Nav1.8 transcript at 7 days (0.76±0.05; P<0.01), 14 days (0.71±0.07; P<0.01) and 21 days (0.76±0.02; P<0.001) and a substantial downregulation of Nav1.9 transcript at 14 days (0.81±0.04; P<0.01) and 21 days (0.806±0.02; P<0.001) in L4–6 ipsilateral DRGs (Fig. 3B); however, no significant differences were observed among groups in the contralateral DRGs (Fig. 3C; P>0.05). Representative images of Nav1.8 and Nav1.9 protein expression within the ipsilateral and contralateral DRGs are shown in Fig. 3D. Consistent with these findings, RT-qPCR analysis revealed that the relative ratio of band intensity of Nav1.8 protein in ipsilateral DRGs was significantly reduced at 7 days (0.62±0.04; P<0.001), 14 days (0.51±0.04; P<0.001) and 21 days (0.63±0.04; P<0.001) after CCI treatment, while Nav1.9 protein expression following CCI exhibited a decrease at 14 days (0.88±0.06; P>0.05) and 21 days (0.89±0.06, P>0.05; Fig. 3E). However, the differences were not statistically significant when compared with the sham group. In addition, there were no detectable differences among contralateral DRG neurons from the rats subjected to CCI (Fig. 3F). These results markedly suggested that Nav1.8 and Nav1.9 sodium channels have distinct roles following CCI treatment.

Effect of CCI on TTX-R sodium channels in rat DRG neurons

The small- and medium-diameter DRG neurons (12–25 µm) were selected as the main focus of the present study. Differences in the voltage protocols and pharmacological inhibition by TTX were used to evoke Nav1.8 and Nav1.9 inward currents (Fig. 4A). Following CCI surgery, the sodium current densities mediated by Nav1.8 and Nav1.9 decreased ~50% (Fig. 4B; between −27.10±6.03 and −14.75±4.01 pA/pF; P<0.01) and 18% (Fig. 4B; between −6.90±2.89 and −5.67±2.00 pA/pF; P>0.05), respectively.

Figure 4 Changes in Nav1.8 and Nav1.9 electrophysiological functions following CCI treatment. (A) Nav1.8 current was evoked by depolarizing voltage steps between −50 and +50 mV in 10-mV increments from a holding potential of −100 mV. The current of Nav1.9 was generated by 5-mV progressive steps between −80 and −35 mV from a holding potential of −100 mV. (B) Following CCI surgery, the peak sodium current density mediated by Nav1.8 was significantly decreased by ~50% (P<0.01). (C and D) The average amplitude of the Nav1.8 and Nav1.9 sodium currents as a function of test pulse voltage in DRG neurons exhibited no change. (E) Dynamics investigations revealed that CCI treatment caused a depolarizing shift of 5.3 mV in the activation curve and ~10 mV positive shift of the inactivation curve of Nav1.8. (F) However, no significant difference was identified between injured and control DRG neurons with regard to the biophysical properties of Nav1.9. DRG, dorsal root ganglion; CCI, chronic constriction injury.

Steady-state activation curves were constructed as described from current-voltage curve experiments. The normalized activation curves of Nav1.8 and Nav1.9 were fitted with the Boltzmann function expressed as: G/Gmax = 1/{1+exp[(V−V1/2)/K, where V1/2 is the membrane potential at half-activation and K represents the slope factor. Characterization of activation curves revealed that CCI treatment caused a depolarizing shift of 5.3 mV and 1.44 mV in Nav1.8 (Fig. 4E; V1/2 between −15.84±0.21 and −10.51±0.24 mV) and Nav1.9 (Fig. 4F; between −40.28±1.02 mV and −38.84±2.07 mV) steady-state activation curves, respectively.

The voltage-dependent inactivation of the Nav1.8 current is shown in Fig. 4E. The solid lines were fitted with the Boltzmann equation, I/Imax = 1/{1+exp[(V−V1/2)/K, where V1/2 is the membrane potential when I/Imax = 0.5 and K represents the slope factor. There was a 10-mV depolarizing shift in the midpoint of inactivation (Fig. 4E; between −23.85±0.74 and −33.73±0.91 mV). This change reflected a parallel shift in the availability curve of the current as no change was observed in the slope factor (between −6.82±0.63 mV and −6.64±0.76 mV). However, no significant differences were identified between injured and control DRG neurons with respect to the biophysical properties of Nav1.9 (Fig. 4F).

Discussion

Neuropathic pain originating from pathology within the nervous system is a serious unmet medical concern. Animal models of neuropathic pain, although often unrepresentative, provide important information for understanding the underlying mechanism of neuropathic pain in humans (27). Peripheral nerve injury may result in pain-associated behavior characterized by spontaneous pain, hyperalgesia and allodynia (28). CCI, as a classical neuropathic pain model, is able to induce spontaneous pain and hyperalgesia through noxious thermal and mechanical stimuli (29). In the present study, it has been demonstrated that CCI is able to stably induce the typical features of neuropathic pain. This method exhibited multiple advantages, including a simple surgical procedure with little tissue damage and a high success rate, with evident and stable spontaneous pain following surgery.

The present findings confirmed the results of previous studies investigating the downregulation of the Nav1.8 current and expression in injured neurons (3,9,14). Nav1.8 mRNA transcript and protein levels were significantly reduced in injured DRGs from 7 days after CCI and the current density mediated by the Nav1.8 channel was markedly reduced at 21 days. However, it remains to be elucidated how this contributes to the neuropathic pain phenotype. Recently, emerging evidence has revealed that the increase in Nav1.8 levels and TTX-R current upregulated in adjacent spared uninjured neurons may provide a reasonable explanation for the role of the Nav1.8 channel in neuropathic pain models (12–14). However, in the present study, the current and expression of Nav1.8 were not affected in contralateral uninjured DRGs. This indicated that there were no redistributed compensatory effects of Nav1.8 in the contralateral uninjured DRGs under CCI-induced neuropathic pain, contrasting with previous studies (15).

A potential reason for the downregulation of the Nav1.8 sodium channel is that the surgical procedure caused neuronal damage. During the present study, it was demonstrated that, in ipsilateral DRGs, CCI treatment induced neuronal damage in a time-dependent manner, exhibiting pyknosis and anachromasis of the nuclei as well as necrosis and shrunken cavities in the cell bodies. Similarly to behavioral assessments, the histological morphology of the contralateral DRGs remained unchanged among groups. It was hypothesized that the quantity of normal neurons affects the excitability of afferent neurons. Thus, functional analysis was performed using the patch clamp electrophysiological technique. It was identified that CCI treatment caused a depolarizing shift in the Nav1.8 steady-state activation curve. This finding is supported by a previous study in which Amm VIII, an α-toxin isolated from venom, is able to induce rapid mechanical and thermal pain hypersensitivities by negatively shifting the activation curve (30). The slowly inactivating Nav1.8 current has been observed to be capable of generating repeated action potentials (31). In the present study, a positive shift was also observed in the Nav1.8 activation curve following the induction of CCI. This may induce the fast inactivated state and increase action potential firing rates.

Another neuronal TTX-R channel, Nav1.9, has also been observed to be associated with neuropathic pain. A previous study revealed that Nav1.9 expression decreased ~3.6 fold under sciatic nerve ligation-induced neuropathic pain (15). However, during the present study, CCI-treatment only led to a significant decrease in the level of Nav1.9 mRNA in DRGs ipsilateral to the injury. In addition, it is important to acknowledge that no change in the relative expression of Nav1.9 protein and Nav1.9 current density and dynamics were observed. This was conflicting with the normal behavior observed in the Nav1.9 knockout mouse following CCI treatment (2). The reasons for this discrepancy remain to be elucidated at present and require further study.

In conclusion, the present study provided evidence of a role for Nav1.8 in the pathogenesis of neuropathic pain. Of note, a positive shift of intact Nav1.8 sodium channels in the DRG neurons ipsilateral to the induced injury was likely to have promoted ectopic discharge, rather than a compensatory regulation of contralateral uninjured DRG neurons.

Acknowledgments

The present study was supported by the National Natural Science Foundation of China (grant no. 81073081), the Excellent Talents Plan of Higher Education Institutes in Liaoning province (grant no. LJQ2013105) and the Key Laboratory of Cardiovascular Medicine Research (Harbin Medical University), Ministry of Education.

References