MiR-30b-5p attenuates oxaliplatin-induced peripheral neuropathic pain through the voltage-gated sodium channel Nav1.6 in rats

Neuropharmacology 153 (2019) 111–120

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MiR-30b-5p attenuates oxaliplatin-induced peripheral neuropathic pain through the voltage-gated sodium channel Nav1.6 in rats

T

Lei Lia,1, Jinping Shaoa,1, Jiangshuan Wangc,1, Yaping Liua, Yidan Zhanga, Mengya Zhanga, Jingjing Zhanga, Xiuhua Rena, Songxue Sua, Yunqing Lid, Jing Caoa,b,∗, Weidong Zanga,b,∗∗ a

Department of Anatomy, School of Basic Medical Sciences, Zhengzhou University, 45001, Zhengzhou, Henan, China Neuroscience Research Institute, School of Basic Medical Sciences, Zhengzhou University, 45001, Zhengzhou, Henan, China c Department of Anatomy, Luohe Medical College, 462000, Luohe, Henan, China d Department of Anatomy, The Fourth Military Medical University, 710000, Xi'an, Shaan Xi, China b

HIGHLIGHTS

1.6 expression increased in the DRGs of oxaliplatin-induced neuropathic pain. • Na expression decreased in the DRGs of oxaliplatin-induced neuropathic pain. • miR-30b attenuated pain by negatively regulating the expression levels of Na 1.6. • miR-30b • miR-30b inhibited the proliferation of colorectal cancer cells. v

v

ARTICLE INFO

ABSTRACT

Keywords: Nav1.6 miR-30b Neuropathic pain Oxaliplatin

Oxaliplatin is a third-generation derivative of platinum that is effective in the treatment of multiple solid tumors. However, it can cause peripheral neuropathic pain, and the molecular mechanisms of this effect remain unknown. We induced a model of peripheral neuropathic pain in rats by intraperitoneally injecting them with oxaliplatin twice a week for 4.5 weeks. We found that both the mRNA and protein expression levels of Nav1.6 (encoded by the gene Scn8a) increased while the miR-30b-5p (shorthand for miR-30b) expression decreased in the dorsal root ganglion (DRG) of treated rats. Using TargetScan and miRanda predictive software, we discovered that Scn8a was a major target of miR-30b. Moreover, we found that miR-30b negatively regulated Scn8a by binding to the Scn8a 3′UTR in PC12 cells. In addition, Nav1.6 and miR-30b were colocalized in the DRG neurons of naive rats. Overexpression of miR-30b using an miR-30b agomir attenuated neuropathic pain induced by oxaliplatin and inhibited both the mRNA and protein expression levels of Nav1.6 both in vitro and in vivo. Conversely, the inhibition of miR-30b with an miR-30b antagomir resulted in neuropathic pain and an increase in the expression of Nav1.6. More importantly, overexpression of miR-30b inhibited the proliferation of LS-174t cells (Colorectal cancer cells). These data suggest that miR-30b contributes to oxaliplatin-induced chronic neuropathic pain through Nav1.6 downregulation and could be a novel therapeutic target for the treatment of oxaliplatin-induced neuropathic pain as a side effect of chemotherapy in cancer patients.

1. Introduction Oxaliplatin, a third-generation platinum derivative, is a valuable drug widely used in the treatment of various solid tumors. Like many other anticancer agents, oxaliplatin treatment leads to a common side

effect called chemotherapy-induced peripheral neuropathy (CIPN), which is the main reason for the dose-limiting toxicity of this drug (Kim et al., 2015). Over 60% of patients completing oxaliplatin chemotherapy complain of persistent neuropathy, with 20–54% of them suffering from chronic pain that can last for months after the end of the

Abbreviations: DRG, dorsal root ganglion; CIPN, chemotherapy-induced peripheral neuropathy; Nav1.6, voltage-gated sodium channels 1.6; PWL, paw-withdrawal latency; OXA, Oxaliplatin; TTX, tetrodotoxin ∗ Corresponding author. Department of Anatomy, School of Basic Medical Sciences, Zhengzhou University, 45001, Zhengzhou, Henan, China. ∗∗ Corresponding author. Department of Anatomy, School of Basic Medical Sciences, Zhengzhou University, 45001, Zhengzhou, Henan, China. E-mail addresses: [email protected] (J. Cao), [email protected] (W. Zang). 1 These authors contributed equally to this study. https://doi.org/10.1016/j.neuropharm.2019.04.024 Received 13 December 2018; Received in revised form 16 April 2019; Accepted 24 April 2019 Available online 03 May 2019 0028-3908/ © 2019 Elsevier Ltd. All rights reserved.

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treatment (Albers et al., 2014). The major symptoms of CIPN include distal and/or perioral paresthesia and muscle fasciculation, which are triggered or enhanced by cold stimulation (Deuis et al., 2013; Xiao et al., 2012). Unfortunately, CIPN is one of the main reasons for the delay or discontinuation of chemotherapy and is therefore responsible for the negative outcomes of cancer management and the resulting poor quality of life. Although the mechanism of CIPN is still unclear, many possible pathways have been suggested. The expression of inflammatory mediators including cytokines and chemokines (Lees et al., 2017), DNA damage (Kelley et al., 2016), and altered intracellular signaling and mitochondrial function have all been implicated in the development of CIPN(Carozzi et al., 2015; Janes et al., 2013). The lack of clarity regarding the mechanism underlying CIPN has hampered the implementation of effective treatment strategies for its relief or prevention. The dorsal root ganglion (DRG) is the first level of neurons where excitability determines the occurrence and development of neuropathic pain (Guerrero-Alba et al., 2018). The excitability of the DRG neurons has been found to be closely related to changes in membrane potential, which depend on changes in ion-channel function. Therefore, ion channels play a vital role in neuropathic pain. Evidence has been presented that oxaliplatin-induced peripheral neuropathic pain is an axonal channelopathy resulting from the modulation of voltage-gated sodium channels (Navs) in neurons. Among the nine sodium-channel subunits (Nav1.1–Nav1.9) expressed in the DRG, Nav1.6 was reported to play an important role in the pain behavior and abnormal spontaneous neuronal activity observed after nerve injury (Xie et al., 2015; Li et al., 2018). Nav1.6 was also shown to be associated with cold allodynia. Therefore, we hypothesized that Nav1.6 participates in the mechanism underlying oxaliplatin-induced peripheral neuropathic pain. There are currently no specific inhibitors of Nav1.6, so we targeted a regulatory mechanism of Nav1.6 to alleviate the oxaliplatin-induced peripheral neuropathic pain. MicroRNAs (miRNAs) are single-stranded, endogenous, small noncoding RNAs (18–25 nucleotides). They regulate gene expression by recognizing the 3′-untranslated region (UTR) of target messenger RNA (mRNA) in a sequence-specific manner and post-transcriptionally inhibiting protein expression (Bartel, 2009). MiRNAs are involved in the regulation of diverse pathological conditions including cancer and neuropathic pain (Tramullas et al., 2018). Although miRNAs have received much attention as therapeutic targets for both neuropathic pain and cancer, the involvement of miRNAs in oxaliplatin-induced neuropathic pain has not been reported. Studies have shown that the miRNA expression profiles varied extensively with different molecular mechanisms; accordingly, distinct miRNAs contributed to different pain conditions. MiR-30b is a miRNA that has been implicated in neuropathic pain, cancer, and neurodegenerative diseases (Su et al., 2017; Croset et al., 2018; Serafin et al., 2015). In this study, we hypothesized that miR-30b inhibits Nav1.6 expression to attenuate oxiplatin-induced neuropathic pain. We found that miR-30b expression was downregulated in DRG neurons following oxaliplatin administration and caused hypersensitivity to mechanical stimuli, possibly through an effect on Nav1.6. Our findings implied that miR-30b could be a novel treatment for the side effects of chemotherapy by acting as a therapeutic target for oxaliplatin-induced neuropathic pain.

University Animal Care and Use Committee. The stock solution of oxaliplatin (1 mg/mL) was diluted in a 5% glucose solution and injected intraperitoneally. Rats underwent intrathecal catheter implantation for drug delivery in the same manner as previously described (Sakurai et al., 2009). Briefly, under 2% isoflurane-induced anesthesia, a lumbar laminectomy of the L5 vertebra was performed and the dura was cut. A sudden movement of the tail or the hindlimb indicated dura penetration. At the L4/5 segment of the spinal cord, a polyethylene-10 catheter was inserted into the subarachnoid space. An intrathecal catheter was implanted in the lumbar enlargement (close to the L4–5 segments) following the method described by Wu (Wu et al., 2004). Following catheter implantation, the animals were given seven days of recovery prior to sham surgery.

2. Materials and methods

2.3. Cell culture

2.1. Animal model and drug infusion

PC12 cells were cultivated in high-glucose Dulbecco's Modified Eagle Medium (DMEM; Solarbio, Hyclone) and supplemented with 5% fetal bovine serum (FBS, Gibco New York, USA), 5% horse serum (Gibco New York, USA), and 1% antibiotics (Gibco New York, USA). The cells were incubated in a humidified incubator with 5% CO2 at 37 °C. After 24 h in culture, when the PC12 cells reached a confluence of 70–80%, they were considered ready for transfection and further

2.2. Behavioral tests 2.2.1. Mechanical paw-withdrawal threshold The latency of paw withdrawal response to mechanical stimulus was determined using the up–down method, following a previously described procedure (Malmquist et al., 2012). Paw-withdrawal responses to mechanical stimuli were measured using a set of von Frey filaments (Muromachi Kikai, Tokyo, Japan). Twenty-four rats were randomly divided into four groups and detected behavioral at 0 d, 1 d, 2 d, 8 d, 9 d, 11 d, 15 d, 22 d, 29 d. Each rat was enclosed in a plastic box with a metallic mesh floor, and a von Frey filament was applied from underneath the mesh floor to the plantar surface of the hind paw. The weakest force (g) that could induce a withdrawal of the stimulated paw in at least three of five trials was referred to as the paw-withdrawal threshold (Su et al., 2017). These behavioral tests were conducted by an investigator blinded to the treatment of the rats. 2.2.2. Thermal paw-withdrawal latency The sample sizes and time points for the thermal trials were the same as those for the mechanical tests. The thermal paw-withdrawal latency (PWL) was measured in the same manner as described by Malmquist (Hargreaves et al., 1988; Malmquist et al., 2012). The rats were placed in Plexiglas chambers that could be heated by aiming a light beam through the glass plate. Radiant heat was delivered to each hind paw through the glass plate, stimulating the middle of the plantar surface (UGOBASILE S.R.L., ITALY). The beam of light was cut off when the rat lifted its hind paw, and the time between the start of the beam of light and the lifting of the hind paw was defined as the PWL. Each trial was repeated three times at 5-min intervals for each hind paw. A maximum shut-off time of 15 s was utilized to avoid any tissue damage. 2.2.3. Cold paw-withdrawal latency The PWL to noxious cold were measured with a cold plate (Zhongbo China) set at 0 °C, the temperature of which was monitored continuously as described previously (Fan et al., 2014; Xu et al., 2014; Li et al., 2015). Each rat was placed in a Plexiglas chamber on the cold plate, which was set at 0 °C. The length of time between the placement of the hind paw on the plate and the animal lifting its foot (with or without paw licking and flinching) was defined as the PWL. The test was repeated three times at 10-min intervals for each hind paw. A maximum cut-off time of 60 s was enforced to avoid tissue injury of the paw pad.

Male Sprague Dawley rats (180–230 g), with access to food and water ad libitum, were housed individually in a clean and open room maintained at a stable temperature (26 °C) on a 12/12 h light/dark cycle. All experimental procedures followed the guidelines of the National Institutes of Health and were approved by the Zhengzhou 112

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experimentation.

Table 1 Primer sets used for qRT-PCR for rat samples.

2.4. Luciferase assay A dual luciferase reporter assay was performed as outlined previously (Huang et al., 2016). The pmirGLO dual-luciferase vector (pmirGLO Vector), which contained both the firefly and renilla luciferase genes, was purchased from Promega (Madison, WI, USA). The 3′UTR of the Scn8a mRNA (which included the predicted binding sites of miR-30b) was inserted into the 3′UTR region downstream of the firefly luciferase gene of the pmirGLO vector (pmir-GLO-UTR). A sitedirected gene mutagenesis kit (GenePharma, Shanghai, China) was used to construct a mutant vector with a mutated miR-30b binding site (pmirGLO-mUTR) as per the manufacturer's protocol. After cultivation for 24 h, PC12 cells were co-transfected with an miRNA mimic (miR30b agomir; GenePharma, Shanghai, China) at different doses of 10, 50, and 100 pM and with the wild-type luciferase reporter (0.5 mg/mL). Transfection was performed with Invitrogen lipofectamine 3000 (Invitrogen, USA). Then, co-transfection of other miRNAs with wild-type and mutant-type reporter vectors was conducted without serum medium or antibody as per the manufacturer's instructions. Co-transfections of the wild-type or the mutant luciferase reporter with an miRNA inhibitor (miR-30b antagomir) or the scramble RNA control were also conducted in a similar manner. After 6 h, we replaced the culture medium with a high-glucose medium containing only 1% antibiotics and 5% FBS. After another 48 h in culture, we used 1× passive lysis buffer to lyse the transfected cells and 20 mL supernatant was collected to measure luciferase activity using the Dual-Luciferase Reporter Assay System (Promega). The ratio of firefly activity to renilla activity was defined as the relative reporter activity. All experiments were performed in triplicate.

Gene name

Primer sequence

SCN8A

5′-AGTAACCCTCCAGAATGGTCCAA-3′ 5′-GTCTAACCAGTTCCACGGGTCT-3′ 5′-TCG GTG TGA ACG GAT TTG GC-3′ 5′-CCT TCA GGT GAG CCC CAG C-3′ 5′-GCT TCG GCA GCA CAT ATA CTA A-3′ 5′-CGA ATT TGC GTG TCA TCC TT-3′ 5′-CCAGCAACTGTAAACATCCTACAC-3′ 5′-TATGGTTTTGACGACTGTGTGAT-3′

GAPDH U6 MiR-30b

China). A template (2 mL) was used for amplification by real-time qPCR with random hexamers, oligo (dT) primers, or specific RT primers as shown in Table 1. GAPDH and U6 were used as reference genes (internal controls) for normalization of Scn8a and miR-30b, respectively. Each sample was run in triplicate in a 20 mL reaction volume containing 250 nM forward and reverse primers, 10 mL SYBR Green qPCR Master Mix (2×, Rox solution provided; Thermo Scientific Maxima,), and 20 ng total cDNA. For qPCR of the reverse-transcribed miRNA, a miRcute miRNA qPCR Detection Kit (SYBR Green, TIANGEN, Beijing, China) was used. The PCR was conducted in a 7500 Fast Real-Time PCR Detection System (Applied Biosystems, USA) and the ratios of the target mRNA levels to the control mRNA levels were calculated using the 2−ΔΔCt method. 2.7. Western blotting Rats were euthanized with isoflurane at 29 d. To ensure a sufficient amount of protein, two unilateral rat DRG neurons (L4-L6) were pooled together. Based on established protocol, the tissue was homogenized in chilled lysis buffer (10 mM Tris, 5 mM MgCl2, 0.25 mM EGTA, 1 mM EDTA, 1 mM DTT, 40 mM leupeptin, 250 mM sucrose). After centrifugation at 4 °C for 15 min at 1500×g, the supernatant was collected to analyze cytosolic proteins and the pellet was collected to analyze nuclear proteins. The protein content of the samples was measured using the Bio-Rad protein assay (Bio-Rad). The samples were heated at 90 °C for 5 min, 20 mg total protein was separated on a 10% sodium dodecyl sulfate-polyacrylamide electrophoresis (SDS-PAGE) gel, about 2 h and electrophoretically transferred onto a polyvinylidene difluoride (PVDF) membrane. After the membranes were blocked with 3% bovine serum albumin (BSA; Solarbio, Beijing, China) in Tris-buffered saline containing 0.1% Tween-20 (TBST) for 2 h, they were incubated in rabbit anti-Nav1.6 (1:200 dilution; Alomone Labs, Israel) and rabbit anti-β-actin (1:1000 dilution; Zhongshan Jinqiao, China) primary antibodies, and then incubated with primary antibodies overnight under gentle agitation. Then, the membranes were incubated in horseradish peroxidase (HRP)-conjugated anti-rabbit secondary antibody (1:1000 dilution; Jackson, USA) and visualized by Clarity Western ECL Substrate (Bio-Rad). Images were taken using FluorChem E (BioRad, Hercules, CA) and the protein bands were quantified via densitometry using Image Lab software (Bio-Rad). All the cytosolic protein bands were normalized to β-actin.

2.5. Cell culture and transfection Culture and transfection of primary DRG neurons were carried out as described elsewhere (Zhao et al., 2013). Three-week old rats were euthanized with isoflurane. All DRG neurons were collected in cold Neurobasal Medium (Gibco/ThermoFisher Scientific USA) with 10% FBS (JRScientific, Woodland, CA, USA), 100 μg/mL streptomycin, and 100 units/mL penicillin (Quality Biological, Gaithersburg, MD, USA). They were then treated with enzyme solution (1 mg/mL collagenase type I, 5 mg/mL dispase, in Hanks balanced salt solution, excluding Mg2+ and Ca2+(Gibco/Thermo Fisher Scientific). The isolated cells were resuspended in mixed Neurobasal Medium and plated in a six-well plate coated with 50 μg/mL poly-D-lysine purchased from Sigma (St. Louis, MO, USA) with a seeding density of 105 DRG neurons/mL. The cells were incubated at 37 °C, 95% O2, and 5% CO2. One day later, Oxaliplatin was added to each 2 mL well 30 min before the small miRNAs (GenePharma, Shanghai, China) were added. 100 μL Neurobasal Medium was used to dilute 5 μL (20 μM) miR-30b agomir/antagomir or 5 μL negative control (20 μM) for 5 min. One hundred microliters Neurobasal Medium was simultaneously used to dilute 2 μL Lipofectamine 2000 (Invitrogen, Carlsbad, CA) for 5 min, then the two solutions were mixed. After 25 min, the mixture was placed into each 2 mL well and 800 Neurobasal Medium was added. The cells were collected 48 h later for PCR and western-blot examinations.

2.8. Immunofluorescence Rats were perfused with 4% paraformaldehyde after they were anesthetized with isoflurane for the preparation of double-labeled immunohistochemistry, as described previously (Xu et al., 2013; Wang et al., 2013). L5 DRG neurons were removed, post-fixed, and dehydrated before frozen sectioning at 16 μm. The L5 DRG sections were pre-incubated in phosphate-buffered saline (PBS) containing 10% normal goat serum and 0.2% Triton X-100 for 30 min and then incubated with primary antibodies at 4 °C overnight. The primary antibodies were rabbit anti-Nav1.6 (1:150 dilution; Alomone labs, Israel),

2.6. Reverse transcription quantitative polymerase chain reaction (RTqPCR) For RT-qPCR, DRG neurons (L4-L6) were pooled to obtain sufficient RNA. Total RNA was extracted using Trizol reagent (Invitrogen, USA), treated using DNase I (New England Biolabs, Ipswich, MA, USA), and reverse-transcribed with the RevertAid First Strand cDNA Synthesis Kit (Thermo, Waltham, USA). MiRNA was reverse-transcribed with an miRcute miRNA First Strand cDNA Synthesis Kit (TIANGEN, Beijing, 113

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mouse anti-NF200 (1:200 dilution; Abcam, Britain), biotinylated antiIB4 (1:100 dilution; Sigma, USA), mouse anti-CGRP (1:50 dilution; Abcam), and mouse anti-gelsolin (GS; 1:200 dilution; R&D). Then, the sections were washed in PBS and incubated with a secondary antibody at 37 °C for 1 h. Images were captured using Leica DMI4000 fluorescence microscope and equipped with a DFC365FX camera (Leica, Germany). 2.9. In situ hybridization Rats were perfused intracardially with 0.9% NaCl followed by 4% cold, buffered paraformaldehyde. The L5 DRGs were extracted and post-fixed in 4% paraformaldehyde overnight. Then, the tissue was cryoprotected by incubation overnight in 30% sucrose at 4 °C. The rat miR-30b in situ hybridization assay kit was purchased from Boster BioTech (Wuhan, China) and the technique was performed as per the manufacturer's protocol. The unique probe sequence for miR-30b was 5′-AGCTG AGTGT AGGAT GTTTA CA-3'. In brief, each section was incubated in a solution of 30% H2O2 and pure methanol (1:50 H2O2: methanol) at room temperature for 30 min, then washed three times with distilled water. To expose the mRNA, 3% citric acid was added to the sections along with two drops pepsase (per 1 mL citric acid) for 2 min at 37 °C. Then, the sections were washed with PBS three times for 5 min each and finally washed with distilled water. Next, the sections were post-fixed in 1% paraformaldehyde in 0.1 M PBS at room temperature for 10 min and washed again. We incubated each section with 20 μL preliminary hybrid liquid for 4 h at 37 °C, and then with a humidified box of 20 mL 20% glycerin for pre-hybridization. Afterward, 20 μL hybrid liquid was applied and the sections were incubated overnight for hybridization at 37 °C. We then washed each section with 2× SSC twice for 5 min each and with 0.5× SSC and 0.2× SSC for 15 min each post-hybridization. After blocking for 30 min at 37 °C, we incubated the sections with anti-rat biotin digoxin at 37 °C for 2 h. Finally, SABC-FITC/CY3 was used for green/red fluorescent staining of the mRNA and the samples were visualized under a fluorescence microscope.

Fig. 1. Oxaliplatin induces mechanical hypersensitivity and cold allodynia, but not thermal allodynia. Paw-withdrawal responses to (a) mechanical, (b) cold, and (c) thermal stimuli after intraperitoneal injection of oxaliplatin in rats. Thermal allodynia and (d) body weight were not changed in all groups. *P < 0.05, **P < 0.01, and ***P < 0.001 vs. the corresponding time points in the control group. n = 6 rats/group.

was administered twice weekly for 4.5 weeks, making a total of nine intraperitoneal injections. These dosages of oxaliplatin caused significant mechanical hypersensitivity (Fig. 1a, F(3,136) = 187.62, P < 0.0001) and cold allodynia (Fig. 1b, F(3,128) = 46.90, P < 0.0001) starting from eight days after the first injection. The mechanical hypersensitivity and cold allodynia were dose-dependent. However, thermal allodynia (Fig. 1c) and body weight (Fig. 1d) did not change over the course of the treatment period in all groups. The absence of changes in body weight showed that the adverse gastrointestinal effects of oxaliplatin were minor. All the rats successfully completed the course of treatment, with no fatalities in any group. Taken together, these data showed that the CIPN model induced by oxaliplatin administration was successful.

2.10. Cellular proliferation assay

3.2. The expression of Nav1.6 is upregulated in the DRG after oxaliplatininduced neuropathy

LS-174t cells were seeded in 96-well plates at a density of 5 × 103 cells per well for 24 h before treatment. After 12 h, 24 h, and 48 h of cultivation, cell proliferation was measured by Cell Counting Kit-8 (CCK-8) system (Dojindo, Japan) according to the manufacturer's instruction. In brief, 10 μL of CCK-8 solution was added to each well and incubated at 37 °C for 1 h. The absorbance was measured at 450 nm with a microplate spectrophotometer (BioTek, USA). There were triplicates for each group, and the experiments were repeated at least three times.

Bulleyaconitine A (intragastric administration, 0.1 mg/kg), a sodium-channel inhibitor, attenuated oxaliplatin-induced peripheral neuropathic pain (Fig. 2a, F(2,66) = 218.92, P < 0.0001). We assessed Nav1.6 expression levels 22 days after the first oxaliplatin injection. The expression levels of Nav1.6 mRNA (Fig. 2b, F(3,15) = 60.98, P < 0.0001) and protein (Fig. 2c, F(3,15) = 15.00, P = 0.0002) were significantly increased corresponding to the increasing doses of oxaliplatin. In addition, the expression of Nav1.6 (Fig. 2d, F(3,11) = 18.67, P = 0.0006 and Fig. 2e, F(3,11) = 52.54, P < 0.0001) in the DRG was also markedly upregulated within each dose of oxaliplatin over the course of the treatment time line (following the first injection). Immunofluorescent staining also showed a significant increase in the ratio of Nav1.6-positive DRG neurons in oxaliplatin-treated rats (Fig. 2f, P < 0.001). Therefore, the change in Nav1.6 expression was dose-dependent and time-dependent in rats with oxaliplatin-induced peripheral neuropathic pain.

2.11. Statistical analysis The data are presented as means ± SEM. For comparisons between two groups, the P value was evaluated and calculated using a two-tailed paired t-test. When there were multiple factors involved, a two-way analysis of variance (ANOVA) followed by Tukey's Multiple Comparison Test was used; multiple groups were compared using a one-way or twoway ANOVA followed by Tukey's Multiple Comparison Test. Values of P < 0.05 were considered statistically significant.

3.3. MiR-30b directly targeted the 3′UTR of Scn8a in PC12 cells

3. Results

The Nav1.6 protein is encoded by Scn8a. We discovered that Scn8a was one of the main targets of miR-30b using TargetScan and miRanda software. The complementary seed sequences (base pair [bp] 53–60) of miR-30b-5p and the Scn8a 3′UTR were conserved between human and rats (Fig. 3a). The expression of miR-30b was significantly downregulated in a time-dependent (Fig. 3b, F(3,11) = 33.29, P < 0.0001)

3.1. Mechanical hypersensitivity and cold allodynia were induced by oxaliplatin treatment We simulated the clinical course of oxaliplatin treatment with different concentrations; 2.4 mg/kg, 3.2 mg/kg or 4.0 mg/kg oxaliplatin 114

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Fig. 2. The expression of Nav1.6 increased in the DRG of rats with oxaliplatin-induced peripheral neuropathic pain. Oxaliplatin was injected into rats at a dose of 4 mg/kg. (a) The paw-withdrawal threshold of Bulleyaconitine A-treated rats was significantly higher than those treated only with oxaliplatin (OXA). **P < 0.01 and ***P < 0.001 vs. OXA group. n = 6 rats/group. The arrows along the X-axis show the times of Bulleyaconitine A treatments. (b and d) Nav1.6 mRNA and (c and e) protein levels were dose-dependent and time-dependent in rats with oxaliplatin-induced peripheral neuropathic pain. *P < 0.05, **P < 0.01, and ***P < 0.001 vs. naïve; #P < 0.05, ##P < 0.01, and ###P < 0.001 vs. day 0. n = 3 rats/group. (f) The ratio of Nav1.6-positive neurons was significantly enhanced by OXA treatment. ***P < 0.001 vs. naïve, Scale bar: 100 μm.

and dose-dependent (Fig. 3c, F(3,11) = 18.50, P = 0.0006)manner in rats with oxaliplatin-induced peripheral neuropathic pain. To determine whether miR-30b actually targeted the 3′UTR sequence of Scn8a mRNA, we inserted the Scn8a 3′UTR sequence downstream of the luciferase gene in a reporter plasmid and measured luciferase activity in the presence of miR-30b. The luciferase signal of the Scn8a 3′UTR plasmid was dose-dependently decreased by miR-30b (Fig. 3d, F(8,35) = 5.57, P = 0.0003). However, the luciferase activity in the presence of an miR-30b antagomir and the scrambled miRNA control was unchanged (Fig. 3e, F(3,11) = 22.63, P = 0.0003). To further prove the specificity of miR-30b and Scn8a binding, we co-transfected a mutant Scn8a 3′UTR luciferase plasmid with the miR-30b agomir into PC12 cells. The luciferase activity of the mutated 3′UTR sequence was not affected by miR-30b (Fig. 3e).

3.4. The localization of Nav1.6 and miR-30b in DRG neurons To examine the localization of Nav1.6 and miR-30b in DRG neurons, we performed double-labeled immunofluorescence and in situ hybridization. As shown in Fig. 4, we co-stained for Nav1.6, the glial cell marker GS (Fig. 4a–c), CGRP that labels small nociceptive peptidergic neurons (Fig. 4d–f), IB4 that labels a subpopulation of small, nonmyelinated nociceptive neurons (Fig. 4g–i), and NF-200 that labels large myelinated non-nociceptive neurons (Fig. 4j–l). The results showed that the Nav1.6 signal was mainly double-labeled with IB4, NF200, and CGRP while it was not found to colocalize with GS. As shown in Fig. 5, the in situ hybridization results revealed that miR-30b was double-labeled with CGRP, IB4 and NF-200 (Fig. 5c, f and i). Importantly, DRG neurons containing miR-30b also expressed Nav1.6 Figure 3. miR-30b directly targeted Scn8a 3′UTR. (a) The matched seed region between miR-30b and the Scn8a 3′UTR predicted by TargetScan is in red. Has, Homo sapien; Rno, Rattus norvegicus. (b) The expression of miR-30b was time-dependently decreased in the DRG of oxaliplatin-treated rats. *P < 0.05 and ***P < 0.001 vs. day 0. (c) The expression of miR-30b was dose-dependently decreased in the DRG of oxaliplatin-treated rats. *P < 0.05 and ***P < 0.001 vs. naive. (d) The miR-30b agomir decreased relative luciferase activity in a dose-dependent manner when co-transfected with the wild-type (WT) Scn8a 3′UTR plasmid in PC12 cells. *P < 0.05 and ***P < 0.001 vs. scramble miRNA. (e) Co-transfection of the miR-30b agomir with WT Scn3a 3′UTR reduced relative luciferase activity, but no change in luciferase activity was detected in the other groups. **P < 0.01 vs. WT + scramble. 115

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(Fig. 5j–l), indicating a potential interaction between miR-30b and Nav1.6. 3.5. Effects of oxaliplatin treatment and miR-30b overexpression or blockade on Nav1.6 expression in primary DRG neurons To verify the effect of oxaliplatin treatment in primary DRG neurons, we added different concentrations of oxaliplatin (0.1, 0.2, and 0.3 μM) into the culture medium and measured the expression of Nav1.6 and miR-30b. We found that all 3 concentrations were able to increase both Scn8a mRNA (Fig. 6a, F(4,14) = 26.95, P < 0.0001) and Nav1.6 protein (Fig. 6b, F(4,19) = 57.93, P < 0.0001) level, while miR-30b was obviously decreased (Fig. 6c, F(4,14) = 14.60, P = 0.0004). Considering drug feature of toxicity, the lowest concertation was applied in following studies. Next, we want to determine whether miR-30b regulates the expression of Nav1.6. Oxaliplatin (0.1 μM) was added to stimulate primary DRG neurons, followed by miR-30b agomir or scramble treatment. The level of miR-30b mRNA was significantly decreased by OXA treatment while it could be rescued by miR-30b agomir (Fig. 6d, F(3,11) = 23.60, P = 0.0003). Meanwhile, oxaliplatin significantly increased Scn8a mRNA (Fig. 6e, F(3,11) = 16.71, P = 0.0008) and Nav1.6 protein (Fig. 6f, F(3,11) = 16.38, P = 0.0009) levels. Encouragingly, miR-30b agomir treatment markedly mitigated the OXA-induced increase in Scn8a mRNA (Fig. 6e) and Nav1.6 protein (Fig. 6f). Furthermore, we found that inhibiting miR-30b (Fig. 6g, F(2,8) = 10.64, P = 0.0106) with miR-30b antagomir efficiently decreased DRG miR30b level, along with a marked increase in the expression of Scn8a mRNA (Fig. 6h, F(2,8) = 44.71, P = 0.0002) and Nav1.6 protein (Fig. 6i, F(2,8) = 11.82, P = 0.0083). Taken together, these results identified a role of miR-30b in regulating Nav1.6 expression and that miR-30b suppressed the expression of Scn8a mRNA and Nav1.6 protein in oxaliplatin-stimulated primary DRG neurons.

Fig. 4. Distribution of the Nav1.6 protein in the DRG neurons of oxaliplatintreated rats. Nav1.6 did not colocalize with the glial-cell marker GS (a–c) but was double-stained with (d–f) CGRP, (g–i) IB4 and (j–l) NF-200. Scale bars: 20 μm.

3.6. Intrathecal administration of the miR-30b agomir inhibited the expression of Nav1.6 in DRG neurons and attenuated oxaliplatin-induced neuropathy To evaluate the exact impact of miR-30b on CIPN, we intrathecally delivered the miR-30b agomir to rats with oxaliplatin-induced peripheral neuropathic pain for four days following day 15 after the first oxaliplatin injection. The mechanical hyperalgesia (Fig. 7a, F(3,119) = 173.38, P < 0.0001) and cold allodynia (Fig. 7c, F(3,132) = 75.51, P < 0.0001) caused by oxaliplatin were attenuated from day 2 following the start of the intrathecal agomir treatment but not treatment with scrambled miRNA. Thermal hyperalgesia was unchanged (Fig. 7b) in all cases. The oxaliplatin-induced upregulation of Nav1.6 was significantly suppressed by the miR-30b agomir in DRG neurons but showed no change in the DRGs of scramble miRNA-injected rats (Fig. 7f, F(3,11) = 17.87, P = 0.0007). The miR-30b agomir also reversed the upregulation of Scn8a (Fig. 7e, F(3,11) = 26.36, P = 0.0002) and downregulation of miR-30b (Fig. 7d, F(3,11) = 21.30, P = 0.0004) in DRG neurons. These results validated that over-expression of miR-30b reversed the rise of Nav1.6 in rats with oxaliplatininduced peripheral neuropathic pain at the level of mRNA and protein, leading to a partial mitigation of CPIN.

Fig. 5. Distribution of miR-30b and colocalization with Nav1.6 in the DRG neurons of naïve rats. In situ hybridization of miR-30b and immunofluorescent staining of (a–c) CGRP, (d–f) IB4, and (g–i) NF200. (j–l) miR-30b was colocalized with Nav1.6. Scale bars: 20 μm.

3.7. Intrathecal miR-30b antagomir upregulated the expression of Nav1.6 in naive rats To further investigate the regulation of Nav1.6 by miR-30b, we downregulated miR-30b by intrathecal injection with miR-30b antagomir in naïve rats. We administered the miR-30b antagomir to naive rats for four days and determined their sensitivity to mechanical, cold, and thermal stimuli. We observed that the paw-withdrawal threshold values in response to mechanical and cold stimuli were markedly lower 116

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Fig. 6. miR-30b regulated Nav1.6 expression in primary DRG neurons. The expression of Nav1.6 (a) mRNA and (b) protein were clearly increased when treated with different concentrations of oxaliplatin (0.1, 0.2, 0.3 μM) in primary DRG neurons. **P < 0.01 and ***P < 0.001 vs. naïve group. (c) The expression of miR-30b was obviously decreased when treated with different concentrations of oxaliplatin (0.1 μM, 0.2 μM, 0.3 μM) in primary DRG neurons. **P < 0.01 naïve group. (d) Expression of miR-30b was significantly decreased by OXA treatment, while it was reversed by miR-30b agomir. **P < 0.01 vs. naïve GROUP; ##P < 0.01 vs. OXA group. (e and f) Over-expression of miR-30b (with miR-30b agomir) attenuated the abnormal up-regulation of Scn8a mRNA and Nav1.6 protein. *P < 0.05 and **P < 0.01 vs. naïve group; #P < 0.05 and ##P < 0.01 vs. OXA group. Compared with naïve group, expression of (g) miR-30b was decreased, while (h) Scn8a mRNA and (i) Nav1.6 protein was increased in primary miR-30b antagomir-treated DRG neurons (inhibitor N.C is scramble of antagomir). *P < 0.05 and **P < 0.01 vs. naïve group. n = 3 in each group.

in rats given the miR-30b antagomir than in naive rats injected with scramble miRNA (Fig. 8a, F(2,57) = 30.54, P < 0.0001 and c, F(2,50) = 17.54, P < 0.0001), proving that the miR-30b antagomir induced pain behaviors in naïve rats. Moreover, downregulation of miR30b (Fig. 8d, F(2,8) = 17.07, P = 0.0033) increased Nav1.6 mRNA (Fig. 8e, F(2,8) = 23.54, P = 0.0014) and protein (Fig. 8f, F(2,8) = 9.94, P = 0.0125) levels in DRG neurons of naive rats.

3.8. miR-30b inhibited the proliferation of LS-174t cells (colorectal cancer cells) To determine whether miR-30b perturbs the proliferation effect of cancer cells in alleviating oxaliplatin-induced peripheral neuropathy, we overexpressed miR-30b in LS-174t human colorectal cancer cells by miR-30b agomir transfection. A cell counting kit 8 (CCK-8) assay showed that overexpression of miR-30b dramatically decreased the growth rate of LS-174t cells compared with scramble-transfected cells (Fig. 9a, F(2,18) = 8.18, P = 0.003). We also suppressed miR-30b Fig. 7. The miR-30b agomir downregulated Nav1.6 and alleviated chemotherapy-induced peripheral neuropathic pain. Paw-withdrawal thresholds in response to (a) mechanical stimuli, (b) cold stimuli, and (c) thermal stimuli after over-expression of miR-30b. *P < 0.05 and **P < 0.01 vs. oxaliplatin group. n = 6 rats/group (d) The relative expression of miR-30b was determined by RT-qPCR in oxaliplatin-injected rats after intrathecal administration of miR-30b agomir. **P < 0.01 vs. oxaliplatin group, ##P < 0.01 vs. naïve group. Nav1.6 (e) mRNA and (f) protein expression were decreased in oxaliplatin-injected rats after intrathecal administration of miR-30b agomir. **P < 0.01 vs. oxaliplatin group. n = 3 rats/group.

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Fig. 8. Inhibition of miR-30b caused Nav1.6 upregulation and induced pain behaviors in naive rats. Paw-withdrawal thresholds in response to (a) mechanical, (b) cold, and (c) thermal stimuli. n = 6 rats/group. The expression of (d) miR-30b and (e) Scn8a mRNA in DRG neurons of naïve rats treated with miR-30b antagomir. (f) The relative protein expression of Nav1.6 in DRG of naïve rats injected miR-30b antagomir. *P < 0.05, **P < 0.01, and ***P < 0.001 vs. naïve group. n = 3 rats/group.

4. Discussion In this study, our data revealed that oxaliplatin induced both cold hyperalgesia and mechanical allodynia in a dose-dependent manner. We observed that the expression of Nav1.6 in DRG was dramatically increased in oxaliplatin-induced CIPN model rats. Meanwhile, enhancing miR-30b significantly reversed the oxaliplatin-induced over-expression of Nav1.6 both in vitro and in vivo. Moreover, intrathecal injection of miR-30b agomir efficiently relieved oxaliplatin-induced neuropathic pain behaviors of CIPN rats. Finally, we revealed that miR30b could inhibit the proliferation of human colorectal cancer cells (LS174t cell line). Oxaliplatin-induced peripheral neuropathic pain exists in almost all patients undergoing chemotherapy. Clinical data demonstrate that the symptoms induced by oxaliplatin can be triggered or enhanced by cold stimulation, and that mechanical and cold pain thresholds decrease progressively with increasing doses of oxaliplatin (Ito et al., 2017; Pachman et al., 2015; De Andrade et al., 2017). In addition, the symptoms can be reported during or immediately after the drug infusion and, in most cases, may last for several months (Janes et al., 2015). Animal models of CIPN using intraperitoneal injections of oxaliplatin have been widely studied, but we simulated the time course of clinical treatment in our animal model of CIPN. Consistent with the clinical data, the dose-dependent mechanical hypersensitivity and cold allodynia induced by oxaliplatin in our CIPN-model rats began between day 8 and day 29 in the treating course (Fig. 1a,c). DRG consist of primary neurons for pain sensation, whose excitability is closely correlated with the resting membrane potential and generation of action potential. The membrane potential of a neuron depends on ion-channel function; therefore, the ion channels in primary DRG neurons determines the occurrence and development of neuropathic pain (Hains et al., 2005; Yang et al., 2018). Some studies showed that the administration of Ca2+ channels antagonist induced an alleviation in paclitaxel-induced neuropathic pain and vincristine-evoked pain. The voltage-gated sodium channel is another important cation channel family whose expression and function directly relate to the generation and development of pain. Nav1.7 expression and function is significantly increased in paclitaxel-induced pain, inflammatory pain, diabetic pain and neuropathic pain (Hong et al., 2004; Chattopadhyay

Fig. 9. miR-30b inhibited the proliferation of LS-174t cells. (a) LS-174t cell proliferation was inhibited by overexpression of miR-30b (agomir transfection) and (b) propagated by the blockade of miR-30b (antagomir transfection). (c) Protein levels of caspase-3 in LS-174t cells were increased after transfection with the miR-30b agomir but (d) decreased after transfection with the miR-30b antagomir. *P < 0.05 and **P < 0.01 vs. control group.

expression in LS-174t cells by transfecting them with the miR-30b antagomir. Blockade of miR-30b markedly increased the growth rate of LS-174t cells compared with cells transfected with scramble miRNA (Fig. 9b, F(2,18) = 7.37, P = 0.0046). Moreover, the protein level of caspase-3 (an apoptosis marker) in LS-174t cells was upregulated after transfection with the miR-30b agomir (Fig. 9c, F(2,8) = 18.67, P = 0.0027) and was downregulated after transfection with the miR30b antagomir (Fig. 9d, F(2,8) = 16.24, P = 0.0038).

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et al., 2008, Huang et al., 2014, Yan Li et al., 2018). Expression of Nav1.3 is also significantly increased in spinal cord injury and SNL models (Su et al., 2017). Besides,Nav1.9 is involved in inflammatory pain (F.R. Nietonet al. 2008). In contrast, the functional roles of Nav1.1 in peripheral sensory neurons is less clear. Our data show that intragastric administration of the sodium channel inhibitor, Bulleyaconitine A, significantly attenuated oxaliplatin-induced peripheral neuropathic pain in rats (Fig. 2a). The result suggests that sodium channels play an important role in CIPN. Several studies have shown that regulation of Nav1.6 contribute to neuropathic pain; the expression of Nav1.6 is altered by spinal nerve ligation and spared nerve injury; and knockdown of Nav1.6 markedly reduced mechanical pain behaviors induced by spinal nerve ligation (Deuis et al., 2013; Patel et al., 2016; Xie et al., 2015; Sittl et al., 2012). Consistent with these studies, we found that suppressing Nav1.6 alleviated oxaliplatin-induced neuropathic pain in rats. Increasing evidence suggests ions channels play an important role in CIPN. The protein and mRNA levels of Nav1.6 were remarkably upregulated in the DRG of oxaliplatin-treated rats. Nav1.6 is a tetrodotoxin (TTX)-sensitive sodium channel expressed in different types of neurons (Deuis et al., 2017), as being confirmed in our study (Fig. 4). Specifically, we observed that Nav1.6 was primarily expressed in medium and large DRG neurons, confirmed by colocalization with NF200 and IB4. These results implied that both myelinated Aβ and Aδ neurons, as well as unmyelinated C neurons in the DRG, were involved in oxaliplatininduced sensory dysfunction. In future experiments, we will use this preparation to investigate the change in the sodium-channel current in an oxaliplatin-induced CIPN model in rats. In recent years, non-coding RNAs have been widely researched as critical regulators of numerous cellular processes involved in disease onset, progression, and prognosis (Lutz et al., 2014; Sakai and Suzuki, 2014). MiRNAs exist extensively in vivo and post-transcriptionally regulate gene expression by repressing mRNA translation or mediating mRNA and protein degradation. There has also been a focus on research linking miRNAs with chronic neuropathic pain states (Lutz et al., 2014). These studies provide foundation for identifying major participating miRNA in oxaliplatin-induced neuropathy. By TargetScan software, miR-30b was predicted to closely target Scn8a. Moreover, we found that miR-30b negatively regulated Scn8a by co-transfecting miR-30b and the Scn8a 3′UTR in PC12 cells (Fig. 3). Our immunofluorescence and in situ hybridization experiments showed that miR-30b was significantly downregulated in the CIPN-model rats and that miR-30b co-expressed with Nav1.6 in rat DRGs, providing direct evidence for the interaction between miR-30b and Nav1.6 (Fig. 5). In this study, we evaluated pain behaviors after manipulating miR30b levels and proved that miR-30b could alleviate nociception in oxaliplatin-treated rats by inhibiting Scn8a. After intrathecal administration of the miR-30b agomir in CIPN-model rats, the observed pain-related behaviors were consistently recovered and the increased expression of Nav1.6 was reversed (Fig. 8). In previous studies of our group, miR-30b was demonstrated to alleviate neuropathic pain by regulating Scn9a (Shao et al., 2016) and Scn3a (Su et al., 2017). Interestingly, some studies show that the pain induced by oxaliplatin do not require the presence of Nav1.7 and Nav1.3 sodium channels (Michael S et al., 2014, Jennifer R Deuis et al., 2013). Taken together, we suggest that miRNAs have diverse roles in regulating pathological conditions and multiple genes can be targeted by a single miRNA. Since our accumulated evidence reveal that miR-30b and Nav1.6 are vital players in oxaliplatin-induced neuropathic pain, miR-30b can be a potential drug target for the treatment of neuropathic pain. In previous study, we also found that over-expression of miR-30b promoted cell apoptosis and suppressed proliferation, migration, and invasion of a gastric cancer cell line, small cell lung cancer and prostate cancer (Zhong K et al., 2014, Kao CJ et al., 2014). In the current study, we transfected miR-30b agomir into LS-174t cells and found that overexpression of miR-30b inhibited tumor cell growth (Fig. 9). Taken

together, our results showed that miR-30b not only alleviated chronic pain but also inhibited the proliferation of human colorectal cancer cells. Although the molecular mechanism for miR-30b inhibiting LS174t cell proliferation is unclear, the results point to a new direction for chemotherapy drug cocktails and provide a useful theoretical foundation for the clinical treatment of oxaliplatin-induced neuropathic pain. There are some limitations in our current study. Firstly, we did not elucidate the upstream mechanism of miR-30b. Secondly, we only measured expression of Nav1.6 and it is important to supplement this with data about ion-channel function. Patch clamp recordings to measure changes in sodium-ion electrical signaling is critical for a follow-up study. Despite these shortcomings, our study revealed that miR-30b directly regulated Scn8a and also presented miR-30b as a new potential target for the treatment of oxaliplatin-induced neuropathic pain. 5. Conclusions We found that miR-30b directly targeted the Scn8a 3′UTR both in vitro and in vivo and alleviated the oxaliplatin-induced chronic neuropathic pain by the negative regulation of Nav1.6 expression in DRG neurons. These findings indicated that miR-30b was involved in the regulation of neuropathic pain by targeting Nav1.6, making this specific miRNA a potential therapeutic target for oxaliplatin-induced chronic neuropathic pain. Funding This work was supported by the National Natural Science Foundation of China (grant number 81671071 and 81471144) and the Natural Science Foundation of Henan Province (number 182300410387). References Albers, J.W., Chaudhry, V., Cavaletti, G., Donehower, R.C., 2014 Mar 31. Interventions for preventing neuropathy caused by cisplatin and related compounds. Cochrane Database Syst. Rev. (3), CD005228. Bartel, D.P., 2009. MicroRNAs: target recognition and regulatory functions. Cell 136, 215–233. Carozzi, V.A., Canta, A., Chiorazzi, A., 2015. Chemotherapy-induced peripheral neuropathy: what do we know about mechanisms? Neurosci. Lett. 596, 90–107. Chattopadhyay, M., Mata, M., Fink, D.J., 2008. Continuous delta-opioid receptor activation reduces neuronal voltage-gated sodium channel (NaV1.7) levels through activation of protein kinase C in painful diabetic neuropathy. J. Neurosci. 28, 6652–6658. Croset, M., Pantano, F., Kan, C., Bonnelye, E., Descotes, F., Alix-Panabieres, C., Lecellier, C.H., Bachelier, R., Allioli, N., Hong, S.S., Bartkowiak, K., Pantel, K., Clezardin, P., 2018. miRNA-30 family members inhibit breast cancer invasion, osteomimicry, and bone destruction by directly targeting multiple bone metastasis-associated genes. Cancer Res. 78, 5259–5273. De Andrade, D.C., Jacobsen, T.M., Galhardoni, R., Ferreira, K., Braz, M.P., Scisci, N., Zandonai, A., Teixeira, W., Saragiotto, D.F., Silva, V., Raicher, I., Cury, R.G., Macarenco, R., Otto, H.C., Wilson, I.B.M., Andrade, D.M.A., Zini, M.M., Henrique, C.D.L., Mendes, B.L., Lilian, R.A., Parravano, D., Tizue, F.J., Lefaucheur, J.P., Bouhassira, D., Sobroza, E., Riechelmann, R.P., Hoff, P.M., Valerio, D.S.F., Chile, T., Dale, C.S., Nebuloni, D., Senna, L., Brentani, H., Pagano, R.L., de Souza, A.M., 2017. Pregabalin for the prevention of oxaliplatin-induced painful neuropathy: a randomized, double-blind trial. Oncologist 22, 1105–1154. Deuis, J.R., Zimmermann, K., Romanovsky, A.A., Possani, L.D., Cabot, P.J., Lewis, R.J., Vetter, I., 2013. An animal model of oxaliplatin-induced cold allodynia reveals a crucial role for Nav1.6 in peripheral pain pathways. Pain 154, 1749–1757. Deuis, J.R., Mueller, A., Israel, M.R., Vetter, I., 2017. The pharmacology of voltage-gated sodium channel activators. Neurophrmacology 127, 87–108. Fan, L., Guan, X., Wang, W., Zhao, J.Y., Zhang, H., Tiwari, V., Hoffman, P.N., Li, M., Tao, Y.X., 2014. Impaired neuropathic pain and preserved acute pain in rats overexpressing voltage-gated potassium channel subunit Kv1.2 in primary afferent neurons. Mol. Pain 10, 8. Guerrero-Alba, R., Valdez-Morales, E.E., Jimenez-Vargas, N.N., Bron, R., Poole, D., Reed, D., Castro, J., Campaniello, M., Hughes, P.A., Brierley, S.M., Bunnett, N., Lomax, A.E., Vanner, S., 2018. Co-expression of mu and delta opioid receptors by mouse colonic nociceptors. Br. J. Pharmacol. 175, 2622–2634. Hains, B.C., Saab, C.Y., Waxman, S.G., 2005. Changes in electrophysiological properties and sodium channel Nav1.3 expression in thalamic neurons after spinal cord injury. Brain 128, 2359–2371. Hargreaves, K., Dubner, R., Brown, F., Flores, C., Joris, J., 1988. A new and sensitive

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