# Upregulation of N-Type Voltage-Gated Calcium Channels Induces Neuropathic Pain in Experimental Autoimmune Neuritis

Authors: Mei Mao; Wen Fan; Yan Zheng; Pan Qi; Min Xi; Yuanrong Yao
Journal: Evidence-Based Complementary and Alternative Medicine
 (2022)

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## Abstract

Objective. Guillain–Barré syndrome (GBS) is a common autoimmune disease of the peripheral nervous system, and there is still no effective treatment for GBS. This investigation intends to figure out the effect and mechanism of N-type voltage-gated calcium (Cav2.2) channels on neuropathic pain in GBS. Methods. An experimental autoimmune neuritis (EAN) model was established in Lewis rats induced by myelin P253-78 peptide and complete Freund’s adjuvant. Luxol fast blue (LFB) staining was used for observing the degree of cell infiltration and demyelination in the sciatic nerve of rats, ELISA for detecting IL-6 and TNF-α expression in the serum, qRT-PCR, and Western blot for measuring the expression of iNOS, MCP-1, and Cav2.2 in the sciatic nerve, respectively. Results. EAN led to significant decreases in the mechanical withdrawal threshold, thermal withdrawal threshold, and mechanical hyperalgesia threshold and an increase in the withdrawal threshold to cold stimulation. The serum IL-6 and TNF-α expression was significantly increased, and the mRNA and protein expression of iNOS, MCP-1, and Cav2.2 in the sciatic nerve were significantly increased in the EAN rats. However, silencing Cav2.2 expression could significantly reverse the above EAN-caused results. Conclusion. Silencing Cav2.2 expression can significantly reduce the clinical score, pathological injury, and mechanical allodynia, reducing the release of inflammatory factors, thus improving neuropathic pain in EAN rats.


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## Body

## 1. Introduction

Guillain–Barré syndrome (GBS) is a common autoimmune disease of the peripheral nervous system and is the most common and severe acute paralytic neuropathy, which affects about 100,000 people each year worldwide [1]. GBS can lead to a disorder that causes your immune system to attack peripheral nerves. The pathological features of the disease are characterized by symmetrical motor weakness of the distal extremities, sensory loss, dysfunction of cranial and autonomic nerves, and, in severe cases, involvement of respiratory muscles [2]. Neuronal cell infiltration and severe demyelination can be observed in pathological sections of peripheral nerves in GBS patients, and even axonal injury in severe cases [1]. The common clinical manifestations of GBS are that 1–3 weeks before the onset, patients have a history of prodromal infection of the upper respiratory tract or gastrointestinal tract or a history of vaccination; most patients begin with symmetrical limb weakness and paresthesia, accompanied by reduced or even absent tendon reflexes in the extremities; the condition progresses to the peak of within about 2–4 weeks after onset [3, 4]. Although most GBS patients recover well after plasma exchange, intravenous gamma globulin, symptomatic and supportive treatment, and immunomodulatory treatment, about 20% of patients leave with varying degrees of physical disability, and about 5% of patients die from the disease or its complications [5]. There is therefore an urgent need to find more effective treatments for GBS.Experimental autoimmune neuritis (EAN) is a classic animal model that mimics the occurrence, development, symptoms, signs and pathological process of GBS patients and serves to investigate the pathogenesis of GBS and develop new treatments. Neurophysiological and histopathological features of EAN are similar to those of GBS [6]. The main approach to inducing the EAN model is to immunize Lewis rats using peripheral nerve myelin proteins such as P0 and P2 [7–10]. However, the pathogenesis of EAN is not yet fully understood, which seriously hinders the research progress of GBS treatment.Voltage-dependent calcium channels are the crucial regulators of neurological functions by controlling intracellular calcium concentrations, including neurodevelopment, neural excitability, and synaptic transmission [11]. N-type voltage-gated calcium (Cav2.2) channels are a subtype of voltage-dependent calcium channels, mainly located in the presynaptic membrane. Cav2.2 mediates the rapid influx of calcium ions into synaptic terminals, thus triggering synaptic vesicle exocytosis as well as neurotransmitter release [12, 13]. Besides, Cav2.2 acts as a functional link between action potentials and neurotransmitter release [14]. Studies have reported that inhibition of Cav2.2 can regulate a variety of neuropsychiatric disorders, so Cav2.2 has become a potential therapeutic target for pain, especially neuropathic pain [15]. However, there is no report regarding the effects of Cav2.2 on EAN and EAN-associated neuropathic pain. Therefore, this study aims to investigate the role of Cav2.2 in regulating neuropathic pain in EAN and its molecular mechanism, providing a new therapeutic direction and reference basis for GBS.


## 2. Materials and Methods

### 2.1. Experimental Animals

A total of 30 healthy male SPF-grade Lewis rats weighing 180–220 g were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. All rats were housed in an environment with a temperature of 22°C, relative humidity of 50%, a 12 h light/12 h dark cycle, a free access to water and food. They were adaptively fed for 7 days before the formal experiments. This study was approved by the Animal Ethics Committee of Guizhou Provincial People’s Hospital (2019-002).


### 2.2. Establishment of Experimental Autoimmune Neuritis (EAN) Model

The EAN rat model was constructed as described previously [16]. After the rats were anesthetized, their hindfoot pads were exposed and injected with 200 μL of antigen solution. 250 μg of peripheral Myelin Protein P2 (53–78) (Cat# 81628-50-4, Shanghai Dongcang Biotechnology Co., Ltd, China) emulsified in an equal volume of complete Freund’s adjuvant (CFA, Sigma-Aldrich, USA) containing 1 mg/ml of heat-killed mycobacterium tuberculosis H37RA. Changes in clinical scores and body weight were recorded daily from day 0 (before immunization) to day 18 (after immunization). Clinical symptom scores were rated as 0 (no disease), 1 (paraparesis of the tail), 2 (paraparesis of the hind limbs), 3 (tetra-paresis), 4 (moribund), and 5 (death); rats with intermediate signs were given an intermediate score of 0.5 [17].


### 2.3. Grouping and Administration

The rats were divided into 5 groups (6 rats/group). Then, 200μL of saline was injected into the hindfoot pads of rats in the control group, 200 μL of CFA in the CFA group, and 200 μL of antigen solution in the EAN group. After injection of 200 μL of antigen solution into the hindfoot pad, the EAN-shRNA group was given 5 mg/kg NC shRNA intraperitoneally (once a day, administered continuously) while the EAN-Cav2.2 shRNA (Supplementary table 1) was given 5 mg/kg Cav2.2 shRNA intraperitoneally (once a day) on the 7th day after immunization. Rats were sacrificed on the 18th day after immunization to collect peripheral blood and isolate sciatic nerves.


### 2.4. Luxol Fast Blue (LFB) Staining

Rat sciatic nerve tissues were fixed in 4% paraformaldehyde, dehydrated using graded alcohols, and finally embedded in paraffin. Paraffin tissue sections were pasted on glass slides, and then dried in a 37°C oven. After that, the glass slides were rinsed using 0.01 mol/L PBS and then placed in a luxol fast blue (LFB) solution (Beyotime, China) in a 37°C oven overnight. The next day, the excess stain was rinsed off with 95% alcohol, and the sections were then rinsed in distilled water. On completion of rinsing, the sections were differentiated with 0.05% lithium carbonate for 10 s, and the differentiation was terminated by rinsing in distilled water. Subsequently, continued differentiation in 70% alcohol was carried out until the gray and white matter were distinguished. Finally, the sections were rinsed, dehydrated, cleaned, and mounted. Under a light microscope, blue represented normal myelin, and the mean number of infiltrating inflammatory cells was counted to assess the degree of demyelination of rat sciatic nerves.


### 2.5. Measurement of Mechanical Withdrawal Threshold by Von Frey Test [18]

The rats were placed in a transparent plexiglass box (22 cm × 12 cm × 22 cm) with a wire grid (hole size: 0.5 cm × 0.5 cm) as the bottom. They were habituated for 30 min before the experiment. The middle sole of the operated hindlimb was stimulated vertically with electronic Von Frey, and the force was slowly applied until the rat lifted or licked the foot. This force was the mechanical withdrawal threshold. This experiment was repeated 4 times to avoid or reduce the effect of the previous stimulation on the subsequent stimulation, and the interval between stimulations at the same site was 3 min. The average value of 4 times was taken as the final result.


### 2.6. Assessment of Thermal Withdrawal Latency by Hargreaves Test [19]

The rats were placed in a transparent plexiglass box (22 cm × 12 cm × 22 cm) with a wire grid (hole size: 0.5 cm × 0.5 cm) as the bottom. They were habituated for 30 min before the experiment. Then the rat sole was irradiated with a BME-410A thermal radiation stimulator. The time from the beginning of irradiation to withdrawal response was the latency to thermal stimulation. The cut-off time was set at 60 s to prevent tissue injury. This experiment was repeated 3 times to avoid or reduce the effect of the previous stimulation on the subsequent stimulation, and the interval between stimulations at the same site was 3 min. The average value of 3 times was taken as the final result.


### 2.7. Measurement of Paw Withdrawal Threshold to Cold Stimulation

0.1 mL of acetone was dripped on the heel of the rat, and rapid paw withdrawal or licking paw represented positive responses. This experiment was repeated 3 times on each side. The number of positive responses per minute was recorded, and the mean value of 3 times was taken as the paw withdrawal threshold to cold stimulation.


### 2.8. ELISA

The serum of rats in each group was taken to detect IL-6 and TNF-α expression according to the instructions of IL-6 and TNF-α ELISA reagents (ClusterTech, China).


### 2.9. qRT-PCR

TRIzol reagent (Thermo Fisher Scientific, USA) was utilized to extract total RNA from rat sciatic nerve tissues. Then cDNA was synthesized by reverse transcription according to the instructions of the reverse transcription PCR kit (Takara, Japan), and the synthesized cDNA was tested for concentration and purity. The real-time PCR reagent (Takara, Japan) was then utilized, and the reaction system was: 95°C for 1 min; 35 cycles of 95°C for 40 s, 58°C for 40 s, and 72°C for 45 s; and 72°C for 10 min. Quantitative analysis was carried out using the 2−ΔΔCt method. The primer sequence used is listed in Table 1.Table 1
Primer sequences.
GeneSequenceiNOSF5′-TGACCATCATGGACCACCAC-3′R5′-ACCAGCCAAATCCAGTCTGC-3′MCP-1F5′-CTCGCCTCCAGCATGAAAGT -3′R5′-GGTGACTGGGGCATTGATTG-3′Cav2.2F5′-AGCCCTCAGATCCCAGCA-3′R5′-GCCTCCTTCTTGCCCTCT-3′GAPDHF5′-GTGAAGGTCGGTGTGAACG−3′R5′-CAATCTCCACTTTGCCACTG−3′


### 2.10. Western Blot

The sciatic nerve tissue of rats in each group was cut into pieces and lysed with RIPA buffer (Gibco, USA) for 20 min. The cells were disrupted by sonication in an ice bath, followed by centrifugation (12,000 rpm, 15 min, 4°C). Then the protein was collected and the protein concentration was measured using a BCA kit (Thermo Fisher Scientific, USA). After SDS-PAGE, the protein was transferred to the PVDF membrane. On completion of a 1 h blocking step with 5% skimmed milk at ambient temperature, primary antibodies iNOS (ab49999, Abcam, UK), MCP-1 (ab7202, Abcam, UK), and Cav2.2 (ACC-002, AlomoneLabs, Israel) were served for overnight coincubation with the membrane at 4°C. The next day, after the membrane was rinsed twice, diluted enzyme-labeled secondary antibody was added and incubated with the membrane for 1 h at ambient temperature. Protein levels were analyzed usingβ-actin as an internal reference.


### 2.11. Statistical Analysis

By utilizing the SPSS 22.0, the experimental data were statistically analyzed. AT-test was employed for comparison between groups, and univariate analysis was conducted for comparison among groups. Mean ± standard deviation (mean ± SD) was the final form to express the results, and P<0.05 was the cut-off value indicating a significant difference.


## 2.1. Experimental Animals

A total of 30 healthy male SPF-grade Lewis rats weighing 180–220 g were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. All rats were housed in an environment with a temperature of 22°C, relative humidity of 50%, a 12 h light/12 h dark cycle, a free access to water and food. They were adaptively fed for 7 days before the formal experiments. This study was approved by the Animal Ethics Committee of Guizhou Provincial People’s Hospital (2019-002).


## 2.2. Establishment of Experimental Autoimmune Neuritis (EAN) Model

The EAN rat model was constructed as described previously [16]. After the rats were anesthetized, their hindfoot pads were exposed and injected with 200 μL of antigen solution. 250 μg of peripheral Myelin Protein P2 (53–78) (Cat# 81628-50-4, Shanghai Dongcang Biotechnology Co., Ltd, China) emulsified in an equal volume of complete Freund’s adjuvant (CFA, Sigma-Aldrich, USA) containing 1 mg/ml of heat-killed mycobacterium tuberculosis H37RA. Changes in clinical scores and body weight were recorded daily from day 0 (before immunization) to day 18 (after immunization). Clinical symptom scores were rated as 0 (no disease), 1 (paraparesis of the tail), 2 (paraparesis of the hind limbs), 3 (tetra-paresis), 4 (moribund), and 5 (death); rats with intermediate signs were given an intermediate score of 0.5 [17].


## 2.3. Grouping and Administration

The rats were divided into 5 groups (6 rats/group). Then, 200μL of saline was injected into the hindfoot pads of rats in the control group, 200 μL of CFA in the CFA group, and 200 μL of antigen solution in the EAN group. After injection of 200 μL of antigen solution into the hindfoot pad, the EAN-shRNA group was given 5 mg/kg NC shRNA intraperitoneally (once a day, administered continuously) while the EAN-Cav2.2 shRNA (Supplementary table 1) was given 5 mg/kg Cav2.2 shRNA intraperitoneally (once a day) on the 7th day after immunization. Rats were sacrificed on the 18th day after immunization to collect peripheral blood and isolate sciatic nerves.


## 2.4. Luxol Fast Blue (LFB) Staining

Rat sciatic nerve tissues were fixed in 4% paraformaldehyde, dehydrated using graded alcohols, and finally embedded in paraffin. Paraffin tissue sections were pasted on glass slides, and then dried in a 37°C oven. After that, the glass slides were rinsed using 0.01 mol/L PBS and then placed in a luxol fast blue (LFB) solution (Beyotime, China) in a 37°C oven overnight. The next day, the excess stain was rinsed off with 95% alcohol, and the sections were then rinsed in distilled water. On completion of rinsing, the sections were differentiated with 0.05% lithium carbonate for 10 s, and the differentiation was terminated by rinsing in distilled water. Subsequently, continued differentiation in 70% alcohol was carried out until the gray and white matter were distinguished. Finally, the sections were rinsed, dehydrated, cleaned, and mounted. Under a light microscope, blue represented normal myelin, and the mean number of infiltrating inflammatory cells was counted to assess the degree of demyelination of rat sciatic nerves.


## 2.5. Measurement of Mechanical Withdrawal Threshold by Von Frey Test [18]

The rats were placed in a transparent plexiglass box (22 cm × 12 cm × 22 cm) with a wire grid (hole size: 0.5 cm × 0.5 cm) as the bottom. They were habituated for 30 min before the experiment. The middle sole of the operated hindlimb was stimulated vertically with electronic Von Frey, and the force was slowly applied until the rat lifted or licked the foot. This force was the mechanical withdrawal threshold. This experiment was repeated 4 times to avoid or reduce the effect of the previous stimulation on the subsequent stimulation, and the interval between stimulations at the same site was 3 min. The average value of 4 times was taken as the final result.


## 2.6. Assessment of Thermal Withdrawal Latency by Hargreaves Test [19]

The rats were placed in a transparent plexiglass box (22 cm × 12 cm × 22 cm) with a wire grid (hole size: 0.5 cm × 0.5 cm) as the bottom. They were habituated for 30 min before the experiment. Then the rat sole was irradiated with a BME-410A thermal radiation stimulator. The time from the beginning of irradiation to withdrawal response was the latency to thermal stimulation. The cut-off time was set at 60 s to prevent tissue injury. This experiment was repeated 3 times to avoid or reduce the effect of the previous stimulation on the subsequent stimulation, and the interval between stimulations at the same site was 3 min. The average value of 3 times was taken as the final result.


## 2.7. Measurement of Paw Withdrawal Threshold to Cold Stimulation

0.1 mL of acetone was dripped on the heel of the rat, and rapid paw withdrawal or licking paw represented positive responses. This experiment was repeated 3 times on each side. The number of positive responses per minute was recorded, and the mean value of 3 times was taken as the paw withdrawal threshold to cold stimulation.


## 2.8. ELISA

The serum of rats in each group was taken to detect IL-6 and TNF-α expression according to the instructions of IL-6 and TNF-α ELISA reagents (ClusterTech, China).


## 2.9. qRT-PCR

TRIzol reagent (Thermo Fisher Scientific, USA) was utilized to extract total RNA from rat sciatic nerve tissues. Then cDNA was synthesized by reverse transcription according to the instructions of the reverse transcription PCR kit (Takara, Japan), and the synthesized cDNA was tested for concentration and purity. The real-time PCR reagent (Takara, Japan) was then utilized, and the reaction system was: 95°C for 1 min; 35 cycles of 95°C for 40 s, 58°C for 40 s, and 72°C for 45 s; and 72°C for 10 min. Quantitative analysis was carried out using the 2−ΔΔCt method. The primer sequence used is listed in Table 1.Table 1
Primer sequences.
GeneSequenceiNOSF5′-TGACCATCATGGACCACCAC-3′R5′-ACCAGCCAAATCCAGTCTGC-3′MCP-1F5′-CTCGCCTCCAGCATGAAAGT -3′R5′-GGTGACTGGGGCATTGATTG-3′Cav2.2F5′-AGCCCTCAGATCCCAGCA-3′R5′-GCCTCCTTCTTGCCCTCT-3′GAPDHF5′-GTGAAGGTCGGTGTGAACG−3′R5′-CAATCTCCACTTTGCCACTG−3′


## 2.10. Western Blot

The sciatic nerve tissue of rats in each group was cut into pieces and lysed with RIPA buffer (Gibco, USA) for 20 min. The cells were disrupted by sonication in an ice bath, followed by centrifugation (12,000 rpm, 15 min, 4°C). Then the protein was collected and the protein concentration was measured using a BCA kit (Thermo Fisher Scientific, USA). After SDS-PAGE, the protein was transferred to the PVDF membrane. On completion of a 1 h blocking step with 5% skimmed milk at ambient temperature, primary antibodies iNOS (ab49999, Abcam, UK), MCP-1 (ab7202, Abcam, UK), and Cav2.2 (ACC-002, AlomoneLabs, Israel) were served for overnight coincubation with the membrane at 4°C. The next day, after the membrane was rinsed twice, diluted enzyme-labeled secondary antibody was added and incubated with the membrane for 1 h at ambient temperature. Protein levels were analyzed usingβ-actin as an internal reference.


## 2.11. Statistical Analysis

By utilizing the SPSS 22.0, the experimental data were statistically analyzed. AT-test was employed for comparison between groups, and univariate analysis was conducted for comparison among groups. Mean ± standard deviation (mean ± SD) was the final form to express the results, and P<0.05 was the cut-off value indicating a significant difference.


## 3. Results

### 3.1. Silencing Cav2.2 Expression Significantly Improves Clinical Scores and Sciatic Nerve Tissue Injury in EAN Rats

First, the effect of silencing Cav2.2 expression on EAN rats was investigated. The results regarding clinical score and sciatic nerve tissue showed that, in comparison with the control group, EAN rats were associated with higher clinical scores and lower body weight, which was statistically significant (P<0.01). Besides, in the EAN group, a large number of inflammatory cells infiltrated the sciatic nerve fiber and endoneurial edema occurred, accompanied by myelin swelling, partial demyelination, and increased pathological score. These results indicated successful modeling. There was no significant difference between the control group and the CFA group. Further, in comparison with the EAN group, silencing Cav2.2 expression significantly reduced the clinical score and increased the body weight, while significantly reduced inflammatory cell infiltration in the sciatic nerve tissue, alleviated demyelination and pathological score (P<0.01 vs. Control group, P<0.01 vs. EAN-shRNA group) (Figures 1(a)–1(c)).Figure 1
Effect of Cav2.2 on clinical score and sciatic nerve tissue of rats. (a) Clinical score record of rats during 18 days of immunization; (b) changes in body weight of rats during 18 days of immunization; (c) luxol fast blue (LFB) staining-based observation of the degree of cell infiltration and demyelination in sciatic nerve tissue of rats on the 18th day of immunization. ∗∗P<0.01 vs. control group, ##P<0.01 vs. EAN-shRNA group.
(a)(b)(c)


### 3.2. Silencing Cav2.2 Expression Alleviates Allodynia in Rats

The effect of silencing Cav2.2 expression on neuropathic pain in rats was further investigated by mechanical, thermal and cold stimulation. The results showed no significant differences in allodynia scores among the groups in the first 6 days; however, from the 7th day, compared with the control group, the mechanical withdrawal threshold, thermal withdrawal threshold, and mechanical hyperalgesia threshold were significantly decreased while the withdrawal threshold to cold stimulation was significantly increased in the EAN group, indicating that the rats had allodynia. However, compared with the EAN group, silencing Cav2.2 induced marked upregulation of the mechanical withdrawal threshold, thermal withdrawal stimulation threshold, as well as mechanical hyperalgesia threshold, and marked downregulation of the withdrawal threshold to cold stimulation (P<0.01 vs. Control group, P<0.01 vs. EAN-shRNA group) (Figures 2(a)–2(d)). Collectively, silencing Cav2.2 expression alleviated allodynia in rats.Figure 2
Effect of Cav2.2 on allodynia in rats. (a–d) Measurement of withdrawal threshold to mechanical (a), thermal (b), and cold (c) stimulation, and mechanical hyperalgesia threshold (d) during 18 days of immunization.∗∗P<0.01 vs. Control group, ##P<0.01 vs. EAN-shRNA group.
(a)(b)(c)(d)


### 3.3. Silencing Cav2.2 Expression Decreases Inflammatory Factor Release and May Be Associated with Improved Inflammatory Cell Infiltration

Since the inflammatory response is vital in autoimmune inflammation, we further investigated the effect of silencing Cav2.2 on the inflammatory response in rats. In comparison with the control group, EAN rats showed a significant increase in serum IL-6 and TNF-α expression, and a marked increase in the mRNA and protein expression of iNOS, MCP-1, and Cav2.2 in the sciatic nerve (P<0.01). In comparison with the EAN group, silencing Cav2.2 caused decreases in IL-6 and TNF-α expression in the serum and mRNA and protein expression of iNOS, MCP-1, and Cav2.2 in the sciatic nerve (P<0.05) (Figures 3(a)–3(c)). Collectively, silencing Cav2.2 expression reduced the inflammatory response in EAN rats, which may be related to the improvement of inflammatory cell infiltration.Figure 3
Effect of Cav2.2 on rat inflammatory response. (a) ELISA-based measurement of IL-6 and TNF-α expression in rat serum; (b) qRT-PCR-based measurement of mRNA expression of iNOS, MCP-1, and Cav2.2 in rat sciatic nerve; (c) western blot-based measurement of protein expression of iNOS, MCP-1, and Cav2.2 in rat sciatic nerve. ∗P<0.05 and ∗∗P<0.01 vs. control group, #P<0.05 and ##P<0.01 vs. EAN-shRNA group.
(a)(b)(c)


## 3.1. Silencing Cav2.2 Expression Significantly Improves Clinical Scores and Sciatic Nerve Tissue Injury in EAN Rats

First, the effect of silencing Cav2.2 expression on EAN rats was investigated. The results regarding clinical score and sciatic nerve tissue showed that, in comparison with the control group, EAN rats were associated with higher clinical scores and lower body weight, which was statistically significant (P<0.01). Besides, in the EAN group, a large number of inflammatory cells infiltrated the sciatic nerve fiber and endoneurial edema occurred, accompanied by myelin swelling, partial demyelination, and increased pathological score. These results indicated successful modeling. There was no significant difference between the control group and the CFA group. Further, in comparison with the EAN group, silencing Cav2.2 expression significantly reduced the clinical score and increased the body weight, while significantly reduced inflammatory cell infiltration in the sciatic nerve tissue, alleviated demyelination and pathological score (P<0.01 vs. Control group, P<0.01 vs. EAN-shRNA group) (Figures 1(a)–1(c)).Figure 1
Effect of Cav2.2 on clinical score and sciatic nerve tissue of rats. (a) Clinical score record of rats during 18 days of immunization; (b) changes in body weight of rats during 18 days of immunization; (c) luxol fast blue (LFB) staining-based observation of the degree of cell infiltration and demyelination in sciatic nerve tissue of rats on the 18th day of immunization. ∗∗P<0.01 vs. control group, ##P<0.01 vs. EAN-shRNA group.
(a)(b)(c)


## 3.2. Silencing Cav2.2 Expression Alleviates Allodynia in Rats

The effect of silencing Cav2.2 expression on neuropathic pain in rats was further investigated by mechanical, thermal and cold stimulation. The results showed no significant differences in allodynia scores among the groups in the first 6 days; however, from the 7th day, compared with the control group, the mechanical withdrawal threshold, thermal withdrawal threshold, and mechanical hyperalgesia threshold were significantly decreased while the withdrawal threshold to cold stimulation was significantly increased in the EAN group, indicating that the rats had allodynia. However, compared with the EAN group, silencing Cav2.2 induced marked upregulation of the mechanical withdrawal threshold, thermal withdrawal stimulation threshold, as well as mechanical hyperalgesia threshold, and marked downregulation of the withdrawal threshold to cold stimulation (P<0.01 vs. Control group, P<0.01 vs. EAN-shRNA group) (Figures 2(a)–2(d)). Collectively, silencing Cav2.2 expression alleviated allodynia in rats.Figure 2
Effect of Cav2.2 on allodynia in rats. (a–d) Measurement of withdrawal threshold to mechanical (a), thermal (b), and cold (c) stimulation, and mechanical hyperalgesia threshold (d) during 18 days of immunization.∗∗P<0.01 vs. Control group, ##P<0.01 vs. EAN-shRNA group.
(a)(b)(c)(d)


## 3.3. Silencing Cav2.2 Expression Decreases Inflammatory Factor Release and May Be Associated with Improved Inflammatory Cell Infiltration

Since the inflammatory response is vital in autoimmune inflammation, we further investigated the effect of silencing Cav2.2 on the inflammatory response in rats. In comparison with the control group, EAN rats showed a significant increase in serum IL-6 and TNF-α expression, and a marked increase in the mRNA and protein expression of iNOS, MCP-1, and Cav2.2 in the sciatic nerve (P<0.01). In comparison with the EAN group, silencing Cav2.2 caused decreases in IL-6 and TNF-α expression in the serum and mRNA and protein expression of iNOS, MCP-1, and Cav2.2 in the sciatic nerve (P<0.05) (Figures 3(a)–3(c)). Collectively, silencing Cav2.2 expression reduced the inflammatory response in EAN rats, which may be related to the improvement of inflammatory cell infiltration.Figure 3
Effect of Cav2.2 on rat inflammatory response. (a) ELISA-based measurement of IL-6 and TNF-α expression in rat serum; (b) qRT-PCR-based measurement of mRNA expression of iNOS, MCP-1, and Cav2.2 in rat sciatic nerve; (c) western blot-based measurement of protein expression of iNOS, MCP-1, and Cav2.2 in rat sciatic nerve. ∗P<0.05 and ∗∗P<0.01 vs. control group, #P<0.05 and ##P<0.01 vs. EAN-shRNA group.
(a)(b)(c)


## 4. Discussion

GBS is a major cause of acute paralysis. It is an immune-mediated disease that may be triggered by recent infections and driven by an immune attack against the peripheral nervous system. Despite progress in research on the treatment for GBS, its morbidity and mortality remain high [20]. Further search for effective treatment strategies is therefore needed. EAN is a classic animal model of GBS. In the absence of specific drugs for GBS, some drugs targeting the humoral and cellular components of the immune response have been used to treat EAN in an effort to find new therapeutic alternatives for GBS [21]. EAN can be induced in by immunization with purified peripheral nerve tissue, myelin protein components P0 or P2 or their synthesized peptides in susceptible animals; EAN can also be induced by injection of antigen-specific autoimmune T cells into rats via the tail vein [10]. Lewis rats show the strongest immune response to EAN induced by P2 protein [22]. Therefore, in this experiment, we selected Lewis rats as experimental animals and P253-78 peptides as sensitizing reagents to construct the EAN model. The rats in the EAN group presented with increased clinical scores, decreased body weight, and mechanical allodynia, confirming successful modeling.Voltage-gated calcium channels are transmembrane protein complexes composed of multiple subunits, including the Cav1 subfamily, Cav2 subfamily, and Cav3 subfamily, all of which regulate the function of pain pathways by affecting neuronal excitability and synaptic transmission [23, 24]. Some studies have reported that calcium channels are involved in the regulation of neuropathic pain [15]. The Cav2.2 gene in the Cav2 subfamily mainly distributed in the presynaptic ends of nerve-muscle encodes N-type calcium channels; the opening of the Cav2.2 channels enables calcium ions to flow into the ganglion, resulting in the release of neurotransmitters [25]. Early studies have found that blocking Cav2.2 channels can produce effective and extensive analgesic effects by inhibiting the release of pain transmitters from the dorsal root of the spinal cord [26]. In addition, it was also found that after knockdown of the Ca2+ 2.2 subunit, the body’s response to neuropathic pain was significantly reduced, and intrathecal administration of specific blockers of Cav2.2 channels was able to significantly inhibit allodynia [27]. In the present study, we confirmed that the mechanical withdrawal threshold, thermal withdrawal threshold, as well as mechanical hyperalgesia threshold of rats were significantly increased after silencing Cav2.2 expression, and allodynia was relieved. LFB staining can show whether the myelin sheath is intact, degenerated, necrotic, and repaired under pathological conditions, so it is significant for both histopathological diagnosis and research of nerves. Silencing Cav2.2 expression was also found in this study to significantly reduce clinical scores and increase body weight in rats, while significantly reducing infiltration of inflammatory cells in the sciatic nerve and alleviating demyelination in rats at the peak of the disease.Nitric oxide (NO) is involved in many physiological processes of the human body and is a critical mediator in regulating pain. NO plays a complex and diverse role in the formation and development of hyperalgesia. Studies have revealed that noxious stimulation causes the release of NO from spinal dorsal horn neurons, and NO interacts with a variety of transmitters to strengthen pain signals in the spinal dorsal horn and produce pathological pain [28]. iNOS is a rate-limiting enzyme for NO synthesis. It has been reported that the NOS inhibitor NG-methyl-L-arginine prevents hyperalgesia caused by PGE2, and can reverse the existing hyperalgesia [29]. CCL2 (or monocyte chemoattractant protein-1, MCP-1), a member of the CC subfamily, has chemotactic activity for monocytes and can activate monocytes and macrophages, which in turn regulate the levels of inflammatory factors such as IL-1 and IL-6 [30]. In the present study, it was further found that silencing Cav2.2 expression significantly decreased inflammatory factor release, and iNOS, MCP-1, and Cav2.2 expression levels were also significantly reduced in the rat sciatic nerve. The above results proved that knockdown of the Cav2.2 gene alleviated neuropathic pain in EAN.


## 5. Conclusion

In summary, silencing Cav2.2 expression reduces clinical scores, pathological injury, and mechanical allodynia, but increases body weight. Silencing Cav2.2 can also decrease inflammatory factor release, which may be associated with decreased inflammatory responses and inflammatory cell chemotaxis. Therefore, upregulation of Cav2.2 channels to activate inflammatory responses may be responsible for causing neuropathic pain in EAN rats.



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*Source: 8547095-2022-06-15.xml*