Male Sprague Dawley (SD) rats, weighting 225 g was used in all presented experiments. Rats were kept in the animal room at a temperature of 24 °C, relative humidity of 60% and 12:12-hour light: dark cycle. All the procedures were approved by the animal ethical committee of Jingzhou Hospital Affiliated to Yangtze University (approval number: JZYY-2020142).

The diabetic neuropathy was induced via intraperitoneal (i.p.) injection of streptozotocin (STZ) as described by previous report [18]. STZ was obtained from TOCRIS (No. 1621, Bio-Techne China, Shanghai, China) and dissolved with sodium citrate buffer (10 mM, pH5.5) to make a 50 mg/mL working solution. For model development, rats were fasted for 6 hours and then injected with a single dose of STZ (45 mg/kg) using a 1 mL syringe. For sham group, rats were administrated with sodium citrate buffer. After injection, all rats were given 10% sucrose water in a period of 48 hours to prevent hypoglycaemics.

AccuChek Performa (Roche Diagnostics, Mannheim, Germany) was adopted to measure non-fasting blood glucose of rats. The blood was taken by puncturing the tip of tail with a hypodermic needle and loaded onto the test strip for measurement. The test was conducted before STZ injection and every week from day 21 to day 49 after injection. The body weight was then recorded after blood test.

The allodynia-like response of rats were measured with an electronic von Frey aesthesiometer (No. 76-0976, Harvard Apparatus, Holliston, MA, USA). The rats were placed in the light chamber on the wire mesh stand. Rats were allowed to acclimate for 15 minutes before test. For measurement, the researcher stabbed the central plantar of rats with the supplied tip and lifted until a pain-like response (i.e., withdrawal, and linking) was observed. The withdrawal threshold of mechanical stimulation was shown in the von Frey meter and recorded. For each time point, all the rats were tested for 5 times in a 5 minutes intervals. The researchers were blinded to the grouping information.

The IITC plantar analgesia meter (390G, IITC life science, Woodland Hills, CA, USA) was applied to measure the changes in heat-evoked response before and after STZ injection. Rats were place in the animal enclosure and allowed to acclimate for 15 minutes before test. A beam of light, which may deliver noxious heat, was aimed at the central plantar of rats until a pain-like response was observed (withdrawal or linking), and the paw withdrawal latency was recorded. For each time point, all the rats were tested for 5 times in a 5 minutes intervals. The researchers were blinded to the grouping information.

The quadripolar SCS electrode and Interstim-II Implantable Pulse Generator (IPG) was obtained from Medtronic (Model 8840, Minneapolis, Minnesota, MN, USA) and implanted on rats from all groups, i.e., control group (without STZ injection), the STZ+sham therapy group (n = 6) and STZ+SCS therapy group (n = 6), at week 3 after i.p. injection. Rats were deeply anaesthetized with isoflurane. A partial laminectomy was performed at the vertebrate level of L1. The electrode was then inserted towards T13 vertebrate level to cover the lumbar spinal cord. The lead of electrode was fixed to the muscles with sutures to secure its position. A IPG was then placed subcutaneously near L1 vertebrate and connected to the lead. After implantation, the muscles and the skins were closed separately. To perform SCS, N’Vision clinician programmer (model 8840, Medtronic, Minneapolis, MN, USA) was adopted to control the electrical delivery by the IPG. First of all, motor threshold was determined by gradually increasing the current amplitude (4 Hz at 200 us pulse width) from 0, until reflection was observed in the hind limb or trunk. For SCS, current amplitude at 80% motor threshold, i.e., 50 Hz at 200 us pulse width was continuously delivered for 12 hours daily, from week 4 after STZ injection, to the end of experiments. For control group and sham therapy group, no currents were delivered.

The relative expression level of Tlr4 and NF-$\kappa$B p65 gene were determined using the qPCR method. Total RNA from lumbar spinal cord dorsal horn tissues were isolated with Trizol reagent (No. 15596026, Thermo Fisher Scientific, San Jose, CA, USA). Dissected tissues were lysed in 1 mL Trizol reagent, following by protein and genomic DNA removal with 200 uL chloroform and total RNA precipitation with 500 uL isopropanol. Complementary DNA (cDNA) was synthesized with 1 ug of total RNA from each rat using the PrimeScript RT reagent kit (No. RR037A, Takara, Dalian, China) according to manufacturer’s instruction. For qPCR, cDNA was diluted for 30 folds with perfect diluent (provided by the kit) and then used for amplification with QuantiTect SYBR Green RT-PCR kit (No. 204156, Qiagen Sciences, Germantown, MD, USA) in an ABI 7900HT system (Thermo Fisher Scientific, San Jose, CA, USA). The relative gene expression levels were normalized to ACTB expression using the 2${}^{-\mathrm{\Delta }\mathrm{\Delta }\text{Ct}}$ method.

The protein level of Tlr4, phosphorylated and NF-$\kappa$B p65 was measured by western blotting. Total protein was extracted from lumbar spinal cord dorsal horn tissues using ReadyPrep${}^{\text{TM}}$ protein extraction kit (No. 1632086, Rio-Rad, Hercules, CA, USA). Protein concentration was detected using Enhanced BCA protein assay kit (No. 23225, Beyotime, Shanghai, China). Protein samples (10 ug/lane) was loaded and separated on SDS-PAGE, then transferred onto 0.22 um PVDF membranes (3010040001, Roche, Nottingham, UK). The membrane was blocked with 5% skim milk in TBST (20 mM Tris, pH8.0, 137 mM NaCl and 0.1% Tween-20) and incubated with primary antibodies (1:1000 dilution) against Tlr4 (#14358S, Cell Signaling, Danvers, MA, USA), phosphor-NF-$\kappa$B p65 (S536, #3033, Cell Signaling, Danvers, MA, USA), NF-$\kappa$B p65 (#8242, Cell Signaling, Danvers, MA, USA) and $\beta$-actin (#3700, Cell Signaling, Danvers, MA, USA) for overnight at 4 °C, following by incubation of corresponding secondary antibodies (1:3000 dilution, Cell Signaling, Danvers, MA, USA) for 1 hour at room temperature. All the antibodies were purchased from Cell signaling technology (Danvers, MA, USA). Finally, the target bands were visualized using SuperSignal West Dura (ThermoFisher Scientific, San Jose, CA, USA) and quantified with Image J (NIH, Baltimore, Maryland, USA).

The protein level of proinflammatory cytokines were measured by ELISA. All the ELISA kits, including rat Il1$\beta$ (SRLB00, Bio-techne China, Shanghai, China), Il6 (SR6000B, Bio-techne China, Shanghai, China) and Tnf$\alpha$ (SRTA00, Bio-techne China, Shanghai, China) quantikine ELSIA kits were purchased from R&D systems (Bio-techne China, Shanghai, China). Spinal cord dorsal horn tissues were collected and lysed using RIPA buffer (ab156034, Abcam, Cambridge, MA, USA) supplemented with protease inhibitor cocktails (ab65621, Abcam, Cambridge, MA, USA). The protein concentration was determined as described above. All the samples were then adjusted to the same concentration with RIAP buffer and loaded to the wells of ELISA plates for 1.5 hours at room temperature. After three washes, Il1$\beta$, Il6 or Tnf$\alpha$ antiserum was added and incubated for 30 min, following with another 3 x washes and antibody conjugate incubation for 30 min. Finally, the substrate solution and stop solution were applied to determine the content of cytokines at 570 nm.

For analysis of three groups across time, two-way analysis of variance (ANOVA) followed by multiple comparisons were conducted. For analysis of four groups at a single time point, one-way ANOVA with multiple comparisons were employed. A p-value less than 0.05 was considered statistical significance.

Rats (n = 12) were assessed for the baseline of paw withdrawal threshold (PWT) and paw withdrawal latency (PWL) by electronic von Frey test and Hargreaves heat hyperalgesia test, respectively. Rats were then randomly assigned to control (vehicle treated) or STZ treated groups according to the baseline. As expected, the blood glucose levels were dramatically elevated in STZ group, compared with the control group. The hyperglycemia was observed at week 3 after STZ injection and lasted for at least for 5 weeks (Fig. 1A). Meanwhile, the body weight was gradually gained in the control group, whereas it was not in the STZ group (Fig. 1B). These results indicated the development of type I diabetes in rats after STZ injection. Furthermore, STZ rats have significantly lower PWT and PWL than control rats, from week 4 to week 7 after model development (Fig. 1C,D). Taken together, we successfully developed the rat model of painful diabetic neuropathy.

Fig. 1.

Single injection of STZ developed painful diabetic neuropathy. (A) Blood glucose test demonstrated STZ injected rats had significantly higher glucose levels than control rats from day 21 to day 49 after injection. (B) STZ injection interfered with gain of body weight. (C) STZ treatment induced heat hyperalgesia on rats from 4 weeks after injection. (D) STZ treatment induced mechanical allodynia on rats from 4 weeks after injection. A and B, Student’s t test. C and D, Two-way ANOVA with post hoc test. *, p 0.05; **, p 0.01. n = 6 rats/group. The error bars represent standard deviation.

We then explored if SCS could improve diabetic neuropathy-induced mechanical allodynia and heat hyperalgesia. Rats (n = 18) were assessed for baseline of pain-like response and randomly assigned to three groups, i.e., control group, STZ+sham therapy group and STZ+SCS therapy group. The electrode and IPG implantation were conducted on rats from all groups at day 21 after model development. Then SCS (12 hours daily, 50 Hz, 200 us pulse width) or sham therapy began from day 21 to the end of experiments. As demonstrated in the Fig. 2A,B, SCS did not improve hyperglycemia and increase body weight among STZ treated rats. There was no significant difference in pain hypersensitivity between sham and SCS groups at post-injection day 21 and day 28. However, the therapeutic efficacy of SCS was evident at day 35, day 42 and day 49, as indicated by a significant increase of both PWL and PWT in SCS group, compared with sham group (Fig. 2C,D). These results suggested that SCS was an efficient approach for relieving diabetic neuropathy-induced pain.

Fig. 2.

SCS alleviated pain hypersensitivity without affecting hyperglycemia. (A) Blood glucose levels were not affected by SCS. (B) SCS did not restore body weight gain. (C–D) SCS alleviated mechanical allodynia and heat hyperalgesia after 2 weeks and lasted for at least 3 weeks. Two-way ANOVA with post hoc test. #, p 0.05; ##, p 0.01. n = 6 rats/group. The error bars represent standard deviation.

Neuroinflammation has been closely associated to the development of central sensitization, which contributes to allodynia and hyperalgesia in various model of neuropathic pain [18, 19]. Therefore, the effects of SCS on neuroinflammation were investigated in this study. qPCR was performed to assess the changes of Tlr4 and NF-$\kappa$B p65 genes. After STZ injection, the mRNA levels of both Tlr4 and NF-$\kappa$B p65 were significantly increased in spinal cord dorsal horn, compared with control group (Fig. 3A,B). Notably, SCS for 5 weeks completely suppressed the up-regulation of Tlr4 and NF-$\kappa$B p65 (Fig. 3A,B). There was no significant difference between control and STZ+SCS group, indicative of suppression of Tlr4/NF-$\kappa$B p65 pathway. This finding was further confirmed by the western blotting assay, which demonstrated that STZ and STZ+sham therapy group had significantly higher level of Tlr4, phosphor-NF-$\kappa$B p65 and NF-$\kappa$B p65 proteins than control group, whereas SCS reversed the elevation of Tlr4/NF-$\kappa$B p65 (Fig. 3C–F).

Fig. 3.

SCS suppressed Tlr4/NF-$\kappa$B p65 pathway in spinal cord dorsal horn. (A–B) Compared with control rats, diabetic rats had significantly higher mRNA of Tlr4 and NF-$\kappa$B p65. SCS, but not sham stimulation suppressed STZ induced gene elevation. (C–F) Western blotting confirmed the alterations of Tlr4, phosphor-p65 and NF-$\kappa$B p65 among groups. One-way ANOVA with post hoc test. **, p 0.01, compared with control group; #, p 0.05, compared with STZ+sham group. n = 6 rats/group. The error bars represent standard deviation.

Next, the pro-inflammatory targets of Tlr4/NF-$\kappa$B p65 pathway was assessed. In line with the alterations in Tlr4/NF-$\kappa$B p65 pathway, ELISA showed that STZ also caused significant increase of Il1$\beta$, Il6 and Tnfa (Fig. 4A–C), which play critical roles in central sensitization. SCS for 5 weeks greatly reduced the elevation of these pro-inflammatory cytokines, as compared with STZ or STZ+sham therapy group (Fig. 4A–C). This result consolidated the suppression of neuroinflammation by SCS.

Fig. 4.

SCS reduced pro-inflammatory cytokines in spinal cord dorsal horn. (A–C) ELISA demonstrated that diabetic rats had significantly higher levels of Il1$\beta$, Il6 and Tnf$\alpha$ than control rats. SCS, but not sham stimulation suppressed STZ induced up-regulation of these three cytokines. One-way ANOVA with post hoc test. ***, p 0.001, compared with control group; #, p 0.05, ##, p 0.01, compared with STZ+sham group. n = 6 rats/group. The error bars represent standard deviation.