Effects of botulinum toxin A on endometriosis‑associated pain and its related mechanism.

Endometriosis (EMS) is a common disease in women aged 25‑45 years, and pain is the main clinical symptom. The primary clinical treatment is surgical excision and drug therapy targeting the ectopic lesions, but these have not been very effective. Botulinum neurotoxin serotype A (BTX‑A) has been reported to be useful in the treatment of pain in a variety of diseases. Based on this, the aim of the present study was to explore the therapeutic effect and mechanism of BTX‑A on EMS. A model of nerve injury induced by oxygen glucose deprivation (OGD) was constructed in PC12 cells and EMS mice. Model cells and mice were treated with different concentrations of BTX‑A to observe the changes in pain behavior, to detect cell viability and the secretion of norepinephrine (NE) and methionine enkephalin (M‑EK) in cells and the spinal cord, and to evaluate the expression of apoptosis‑related molecules in spinal cord nerves. The results revealed that BTX‑A significantly reduced the amount of writhing in model mice, enhanced the activity of PC12 OGD cells, increased the secretion of NE and M‑EK in model cells and the spinal cord of mice, and decreased the apoptosis of neural cells in the spinal cord of the model mice. Therefore, it was hypothesized that BTX‑A may alleviate the pain induced by EMS by increasing the secretion of analgesic substances and promoting the repair of nerve injury. The present study provided a theoretical basis for the treatment of pain induced by EMS.

Introduction

Endometriosis (EMS) is a common chronic clinical gynecological disease. It is caused by ectopic activation of endometrial cells and implantation outside of the uterine cavity (1). The incidence of EMS in women aged 25–45 years is ~10% and increases annually (2). At present, the primary treatment methods for the disease are surgical resection of the ectopic endometrium and drug therapy. However, patients are prone to relapse of pain, as well as possible drug addiction or tolerance, therefore, to date there is no effective treatment. Pain is the main clinical symptom of EMS, but the underlying mechanism is not clear (3–5). EMS-induced pain may be caused by the local inflammatory response stimulated by periodic hemorrhage in ectopic lesions, chronic inflammation in the abdominal cavity and the abnormal growth of lesions, which may result in peripheral and central nervous sensitization (6). Prolonged exposure to pathogens or systemic inflammation can lead to chronic, insoluble neuroinflammation, which in turn leads to functional and structural changes until the nerve cells die (7).

Botulinum neurotoxin serotype A (BTX-A) is a neuromuscular toxin secreted by Clostridium botulinum with high toxicity and pathogenicity (8). First, BTX-A binds to the presynaptic nerve endings. Upon binding, BTX-A is transferred to cells through endocytosis, cleaving the related core proteins of the neuroexocytosis apparatus and inhibiting the release of acetylcholine in the synaptic vesicles. Eventually, the signal transduction of neuromuscular junctions is suppressed (8). It is widely used in the treatment of some diseases related to muscle hyperresponsiveness, such as squint, blink, torticollis, hemifacial spasm and cerebral palsy (9,10). At present, BTX-A has been used in the treatment of clinical pathological pain, such as migraine, musculoskeletal pain and refractory trigeminal neuralgia, and has achieved satisfactory efficacy (11,12). In addition, further research has demonstrated that BTX-A can also inhibit the activation of microglia and play a long term role in pain relief in animal models of inflammation and neuropathic pain (13). A previous study has also demonstrated that BTX-A can be used to repair nerve injury and promote the regeneration and growth of neurons (14).

To the best of our knowledge, no previous studies have reported on the application of BTX-A for EMS-associated pain. Therefore, the present study constructed a model of nerve injury induced by oxygen glucose deprivation (OGD) in PC12 cells in vitro and in mice with EMS, in order to investigate whether BTX-A alleviates the pain induced by EMS in mice. These models were then used to study the effects of EMS and OGD on nerve injury repair and neuronal apoptosis, as well as its molecular mechanism in order to provide an experimental basis for the clinical treatment of EMS.

Materials and methods

Cell culture

PC12 cells were purchased from The Cell Bank of Type Culture Collection of the Chinese Academy of Sciences. The cells were seeded in Dulbecco's modified Eagle's medium (DMEM; Gibco; Thermo Fisher Scientific, Inc.) with 10% (v/v) fetal bovine serum (FBS; Gibco; Thermo Fisher Scientific, Inc.), 100 U/ml penicillin and 100 µg/ml streptomycin (Gibco; Thermo Fisher Scientific, Inc.) with a density of 2×106 cells in 10 cm dishes. Cells were maintained at 37°C in a humidified incubator containing 5% CO2. Cell culture medium was replaced every two days.

PC12 OGD model

The PC12 cell suspension was seeded in 96-well culture plates (2×104 cells/well) and incubated at 37°C in a 5% CO2 incubator. After 12 h of normal culture, the cells were cultured in DMEM without glucose in a hypoxic chamber (1% O2, 94% N2, and 5% CO2) for 12 h. After OGD, the culture was washed with PBS three times and then cultured in normal DMEM under normoxic conditions for 24 h. BTX-A (100 U/box; Lanzhou Institute of Biological Products) was separately added during the OGD and reoxygenation culture. Control cells without OGD were maintained under normal conditions. Finally, the culture medium was discarded, cell samples were collected and cytological tests were conducted.

Determination of cell viability

The PC12 cells were collected, and viability was detected by a Cell Counting Kit (CCK)-8 assay with CCK-8 (cat. no. CK04; Dojindo Molecular Technologies, Inc.). According to the kit's protocol, each well was mixed with 10 µl CCK-8, and incubated at 37°C for 1 h. Cell viability was determined using an enzyme-labelled meter (Multiskan™ Mk3; Thermo Fisher Scientific, Inc.) at a wavelength of OD 450 nm. Each experiment was performed in 6 parallel wells and repeated 3 times.

Immunocytochemistry

The PC12 cells (1×104 cells/well), after BTX-A treatment, were fixed in 4% paraformaldehyde at room temperature and then permeabilized in PBS containing 0.5% Triton-X 100 for 15 min at 4°C. Coverslips were incubated in 4% bovine serum albumin (cat. no. 37525; Thermo Fisher Scientific, Inc.) at room temperature for 1 h to block non-specific antibody-binding sites, and then incubated overnight with primary antibodies against β3-tubulin (1:500; cat. no. ab52623; Abcam) at 4°C, followed by incubation with the appropriate Alexa Fluor 594 goat anti-rabbit antibody (1:2,000; cat. no. R37117; Thermo Fisher Scientific, Inc.) for 1 h at room temperature. Then, cells were stained with DAPI for 10 min at room temperature, and examined using a Zeiss LSM710 fluorescence microscope.

Enzyme-linked immunosorbent assay (ELISA)

The levels of norepinephrine (NE) and methionine enkephalin (M-EK) in the PC12 cell supernatant and mouse spinal cord homogenate were detected using ELISA methods, and the operation was performed according to the instructions of the ELISA kit (cat. nos. E-EL-0047c and E-EL-0020c; Elabscience Biotechnology Co., Ltd.). The absorbance was measured at 450 nm using an enzyme-labelled meter (Multiskan Mk3; Thermo Fisher Scientific, Inc.). Each sample was repeated in triplicate.

Mouse EMS model

The present study constructed a mouse model of intraperitoneal EMS. The mice were purchased from Shanghai Sippr-BK laboratory animal Co., Ltd. All mice received food and water ad libitum and were maintained at 26±1°C and 50±5% relative humidity under a 12/12-h light/dark cycle. To establish this model, 6-week-old (18–20 g) female BALB/c mice (n=20) were fed adaptively for 1 week with a standard diet and water ad libitum. The mouse EMS model was established as described previously (15). Briefly, estradiol benzoate at 0.5 mg/kg was administered to donor animals by gavage for 2 days to induce the estrus period. Vaginal smears showed a large number of seedless keratinocytes in mice as donors. After a week, the uteruses of the donor mice were collected, and minced using scissors in sterile normal saline. Then, the uterus tissue debris was injected into the recipient mouse intraperitoneally (i.p.) and half of the preparation was injected into the peritoneum of each of two recipient mice with a syringe. A week later, the model mice were injected intraperitoneally with oxytocin at 20 IU/kg; observing evident pain behavior was defined as successful model mouse establishment (16). Following the removal of unsuccessful model mice, the EMS mice were randomly divided into the following groups: i) Control (n=5), normal mice with saline (i.p.); ii) Sham (n=5), model mice with saline (i.p.); iii) BTX-A high dose (BH, n=5), 30 U/kg (i.p.); and iv) BTX-A low dose (BL, n=5), 10 U/kg (i.p.). Oxytocin 20 IU/kg was injected intraperitoneally at every time point to observe the writhing reaction within 30 min on days 0, 1, 3, 5, 7, 14, 21 and 28 after BTX-A treatment. Among them, oxytocin was injected 1 h after BTX-A treatment, and the time point was defined as 0 days. No model mice died during the behavioral observation period, and there were also no significant weight changes or behavioral variations. After 28 days of observing animal behavior, the mice were sacrificed by cervical dislocation, and spinal cord tissue was collected. The experiments and procedures using mice were approved by the Animal Ethics Committee of Shanghai No. 10 People's Hospital of Tongji University (SHDSYY-2019-T0018) where the experiments were performed.

Writhing test

Writhing test was performed to explore the effect of BTX-A on the pain caused by EMS. Firstly, BTX-A was injected subcutaneously 1 week after the model was established. Then, oxytocin was intraperitoneally injected at 0, 1, 3, 5, 7, 14, 21 and 28 days, respectively. Immediately following the oxytocin injection, visceral pain was measured by counting the number of writhing reflexes for 30 min in the experimental groups. The writhing reflexes were characterized by the presence of abdominal contraction, body distortion, hind limb extension and hip elevation. The investigator was blind to the drug treatment administered. The analgesic response was quantified as the percentage of the number of writhing decreases after treatment with oxytocin, which was calculated as the analgesic frequency. Analgesic frequency=[(number of writhes in sham-number of writhes in determined dose or control)/number of writhes in sham] ×100% (17).

Western blot analysis

Total protein was extracted from tissues using radioimmunoprecipitation assay lysis buffer (cat. no. 87787; Thermo Scientific, Inc.), and the total protein concentration was determined using the BCA assay (cat. no. A53225; Thermo Scientific, Inc.). Protein (20 µg) of each sample was separated via 10 or 12% SDS-PAGE. After SDS-PAGE, proteins were transferred to polyvinylidene fluoride membranes, and prior to incubation with primary antibodies, the membranes were treated with 0.1% Tween-20 in Tris-buffered saline (TBST) containing 50 g/l skimmed milk at room temperature for 4 h. Membranes were incubated with primary antibodies overnight at 4°C and then with a secondary antibody at room temperature for 2 h. The primary antibodies included C-X3-C motif chemokine receptor 1 (CX3CR1; 1:1,000; cat. no. ab8021; Abcam), purinoceptor P2X 7 (P2X7; 1:200; cat. no. sc-514962; Santa Cruz Biotechnology, Inc.), phospho-p38 mitogen-activated protein kinase (MAPK; 1:1,000; cat. no. 4511; CST Biological Reagents Co., Ltd.), Bax (1:3,000; cat. no. ab32503; Abcam), Bcl2 (1:1,000; cat. no. 15071; CST Biological Reagents Co., Ltd.) and β-tubulin (1:500; cat. no. sc-101527; Santa Cruz Biotechnology, Inc.), which was used as the control. Then, the blocked membranes were incubated with rabbit anti-mouse secondary antibody (1:3,000; cat. no. 315-035-003; Jackson ImmunoResearch Laboratories, Inc.) and goat anti-rabbit secondary antibody (1:3,000; cat. no. 111-035-008; Jackson ImmunoResearch Laboratories, Inc.). Bands were detected with a chemiluminescence reagent (cat. no. 34577; Thermo Fisher Scientific, Inc.) and the protein bands were quantified by densitometry using ImageJ version 1.48 (National Institutes of Health).

Statistical analysis

All experiments were repeated at least three times. The data are presented as the mean ± standard deviation of the mean in the graphs. Statistical analysis was performed using paired Student's t-test or one-way analysis of variance followed by Tukey's post hoc test with GraphPad Prism 5 software (GraphPad Software, Inc.). P<0.05 was considered to indicate a statistically significant difference.

Results

BTX-A treatment in vitro can reverse cell damage induced by an OGD model

In order to evaluate the protective effect of BTX-A on PC12 cell injury, according to previous research, the present study established an OGD cell injury model in PC12 cells and β3-tubulin was used as a neuron-specific marker (18–20). By observing the morphology of the cells, the control PC12 cells were reported to be spindle or polygonal in shape. The cells had longer processes and interlaced with each other. After OGD treatment, the cells became round, the processes became shorter, and the cells floated to form clusters. Treatment with two different concentrations of BTX-A gradually reversed the effects observed in OGD cells, and the effect of a high concentration was the most notable (Fig. 1A and B). In addition, CCK-8 was used to detect the effect of BTX-A on the viability of PC12 cells in the OGD model. The results revealed that compared with the control group, the cell viability of the OGD-treated group was significantly decreased (P<0.001) to ~half that of the control group, whereas BTX-A could increase the OGD-induced reduction in cell viability in a concentration-dependent manner (OGD-BL vs. OGD, P<0.001; OGD-BH vs. OGD, P<0.001; Fig. 1C).

Figure 1.

BTX-A treatment can restore the morphology and viability of OGD model cells. (A) Cell morphology after BTX-A treatment (magnification, ×200). (B) Immunofluorescence labeling of cells in each treatment group using the neuronal marker β3-tubulin (magnification, ×200). (C) CCK-8 was used to detect the viability of PC12 OGD model cells treated with different concentrations of BTX-A. ***P<0.001, as indicated. BTX-A, botulinum neurotoxin serotype A; OGD, oxygen glucose deprivation; CCK-8, Cell Counting Kit-8; BL, BTX-A low dose group; BH, BTX-A high dose group.

Activation of BTX-A on neurotransmitter secretion in PC12 cells of OGD

In order to study the possible mechanism of pain relief involved in the repair of nerve cell injury caused by BTX-A, ELISA was performed in the present study to detect the secretion of neurotransmitters in the cell supernatant. The results showed that BTX-A could significantly increase the secretion of NE and M-EK in PC12 cells (Fig. 2A and B). Therefore, the analgesic effect of BTX-A is likely achieved by increasing the analgesic factors secreted by cells.

Figure 2.

Botulinum neurotoxin serotype A increases neurotransmitter secretion in OGD model cells. (A) NE and (B) M-EK. Data are presented as the mean ± standard deviation of three independent biological repeats. *P<0.05, as indicated. OGD, oxygen glucose deprivation; NE, norepinephrine; M-EK, methionine enkephalin; ns, not significant; BL, botulinum neurotoxin serotype A low dose group; BH, botulinum neurotoxin serotype A high dose group.

Effect of BTX-A on pain induced by EMS

The results revealed that the amount of writhing in the control group was almost none within one month. The number of writhing events in the Sham and BTX-A-treated groups were significantly higher than those in the control group (21±1, 22±4 and 26±2 on day 0, respectively). The writhing frequency of the control group and Sham group remained relatively stable at the different time periods, whereas the writhing frequency of the BTX-A group decreased sharply as time increased, and the trend remained stable after the fifth day; the results were slightly higher than those observed in the control group. The trend in writhing times in the high and low BTX-A dose groups were very similar within the one month period, and there was no significant difference between the two groups (Fig. 3A and Table I). In general, BTX-A could relieve the pain induced by EMS.

Figure 3.

Statistical analysis of number of writhes at 0, 1, 3, 5, 7, 14, 21 and 28 days post-surgery. The control group consisted of normal mice given saline, the Sham group was treated with saline after the uterine heterotopia model establishment, and the BTX-A group was treated with BTX-A at the corresponding concentration after uterine heterotopic model establishment. Data are presented as the mean ± standard deviation. Sham group vs. Control group, ***P<0.001; BL group vs. Sham group, ##P<0.01 and ###P<0.001; BH group vs. Sham group, ΔP<0.05 and ΔΔP<0.01 and ΔΔΔP<0.001. BTX-A, botulinum neurotoxin serotype A; BL, BTX-A low dose group; BH, BTX-A high dose group.

Table I.

Establishment of mouse endometriosis model and detection of pain behavior. Data are presented as the mean ± standard deviation.

A, Day 0
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	1±0.57	96.8%
Saline	5	21±1	–
10 U/kg BTX-A	5	22±4	−5.6%
30 U/kg BTX-A	5	26±2	−24.8%
B, Day 1
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	1±1.15	97.1%
Saline	5	23±3	–
10 U/kg BTX-A	5	16±3	29.7%
30 U/kg BTX-A	5	21±3.6	10%
C, Day 3
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	0±0	100%
Saline	5	27±2	–
10 U/kg BTX-A	5	10±3	61.75%
30 U/kg BTX-A	5	12±2	55%
D, Day 5
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	1±1	94%
Saline	5	26±2	–
10 U/kg BTX-A	5	4±3	85.3%
30 U/kg BTX-A	5	5±1	80.7%
E, Day 7
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	0±0	100%
Saline	5	25±2	–
10 U/kg BTX-A	5	3±2	89.7%
30 U/kg BTX-A	5	6±3.21	77.6%
F, Day 14
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	0±0.57	98.5%
Saline	5	22±3	–
10 U/kg BTX-A	5	6±3	74.9%
30 U/kg BTX-A	5	6±2.88	71.6%
G, Day 21
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	2±2.08	92%
Saline	5	21±2	–
10 U/kg BTX-A	5	9±4	58%
30 U/kg BTX-A	5	5±5.13	74.6%
H, Day 28
Groups	Samples	Writhing frequency	Analgesic rate
Control	5	0±0	100%
Saline	5	23±2	–
10 U/kg BTX-A	5	5±2	77.9%
30 U/kg BTX-A	5	6±4.04	72%

BTX-A, botulinum neurotoxin serotype A.

Protective capability of BTX-A on spinal cord neurons in mice

As EMS induces inflammatory infiltration around the visceral tissue of the ectopic site, inflammatory factors constantly stimulate nearby neurons. Long-term stimulation can cause neuronal damage, and nerve injury may lead to long-term pain (7). Next, the present study investigated whether the pain relief induced by BTX-A in EMS was achieved by repairing spinal cord neuron injury. The results showed that the amount of β3-tubulin was notably lower in the Sham group when compared with the control group, but increased to a certain extent in the BTX-A treatment group (Fig. 4). It was suggested that BTX-A may repair spinal cord neuron injury to some extent in vivo.

Figure 4.

Effect of botulinum neurotoxin serotype A on spinal cord neurons in mice. Spinal cord tissues were labeled with β3-tubulin (magnification, ×400); There were 5 rats in each treatment group, and at least three sections were labeled with antibodies in each group. BL, botulinum neurotoxin serotype A low dose group; BH, botulinum neurotoxin serotype A high dose group.

Effect of BTX-A on neurotransmitter secretion in the spinal cord neurons of mice

The expression of NE and M-EK in the spinal cord homogenate of each treatment group was analyzed. The results showed that the expression of NE and M-EK in the Sham group was not changed when compared with the control group, but the BTX-A treatment groups showed different degrees of increase. In the 30 U/kg BTX-A treatment there was a significant difference compared with the Sham group (Fig. 5). From the aforementioned results, it can be concluded that BTX-A can increase the release of NE and M-EK in the spinal cord of mice with EMS. The pain relief induced by NTX-A is likely due to an increase in neurotransmitter secretion.

Figure 5.

Effect of BTX-A on the secretion of spinal cord analgesic factors. BTX-A can increase the secretion of spinal cord analgesic factors (A) NE and (B) M-EK in endometriosis model mice (n=5 rats/group). *P<0.05 and ***P<0.001, as indicated. BTX-A, botulinum neurotoxin serotype A; NE, norepinephrine; M-EK, methionine enkephalin; ns, not significant; BL, BTX-A low dose group; BH, BTX-A high dose group; ns, not significant.

Effect of BTX-A on the apoptosis of spinal cord neurons in mice

A previous study demonstrated that the MAPK family member p38 MAPK signaling pathway is closely related to nerve injury and regulates numerous physiological processes, such as cell differentiation, cell growth and apoptosis. Among them, the ratio of Bcl2/Bax is an important component and an important index to determine apoptosis (21). Therefore, the present study further investigated the effect of BTX-A on neuronal sensitization and apoptosis. Western blotting results showed that the microglia receptor CX3CR1 and P2X7 of chemokines fractalkine increased in the sham group compared with the control group. The results of the present study suggested that BTX-A may induce neuronal sensitization and injury by stimulating the activation of microglia around the spinal neurons and releasing a large number of inflammatory factors, or may induce self-sensitization via the activation of P2X7 on the surface of spinal neurons. Further research is required for confirmation. Similarly, the levels of phosphorylated p38 MAPK and Bax increased, whereas the expression of Bcl2 decreased in the EMS model, but following BTX-A treatment expression levels were close to those of the control group (Fig. 6A and B).

Figure 6.

Effect of BTX-A on the apoptosis of spinal cord neurons in mice. (A) Representative western blotting images of corresponding protein expression in model mice treated with BTX-A. (B) Protein levels were semi-quantified using ImageJ software. Data are presented as the mean ± standard deviation. ns=no significance, *P<0.05 and **P<0.01, as indicated. β-tubulin was used as the internal loading control. CX3CR1, C-X3-C motif chemokine receptor 1; P2X7, P2X purinoceptor 7; p-, phosphorylated; MAPK, mitogen-activated protein kinase; BTX-A, botulinum neurotoxin serotype A; BL, BTX-A low dose group; BH, BTX-A high dose group.

Discussion

PC12 is a rat adrenal pheochromocytoma cell line with some characteristics of adrenal medullary chromaffin cells (18). As adrenomedullary cells originate from the embryonic nerve spine, PC12 cells have neuronal cellular properties and in conventional culture they secrete catecholamines (dopamine, NE and norepinephrine) and enkephalin neurotransmitters, which are analgesic substances (22,23). Therefore, the OGD model was used in the present study to produce neuronal damage to investigate the molecular mechanism underlying the effect of BTX-A on neuronal injury and pain in vitro. The results revealed that BTX-A treatment could significantly increase the survival rate of PC12 OGD model cells and restore the morphology of the cells close to that of the control group in a concentration-dependent manner. This result was similar to previous research, namely, that BTX-A can induce nerve outgrowth (14) and enhance Schwann cell proliferation (24). At the same time, the repair of injured neurons was accompanied by an increase in the secretion of the analgesic factors NE and M-EK. Previous studies have reported that NE and M-EK have analgesic effects. Injection of NE and/or opioid agonists into the spinal dorsal horn can achieve analgesic effects (25,26). Microencapsulated PC12 cells can be transplanted into the subarachnoid space of rats, and as a source of microcellular pump, can continuously produce NE and M-EK, and may have a notable analgesic effect in a chronic neurogenic animal model (23,27). NE is important in the inhibition of neuropathic pain in the spinal cord, primarily via α2-adrenergic receptors, and it also improves the function of the descending noradrenergic inhibitory system (28,29). M-EK, through its receptor, regulates the ion channels of neurons and inhibits the release of excitatory transmitters (30). According to the in vitro results of the present study, BTX-A can repair the PC12 cell damage induced by OGD to some extent, and may have an analgesic effect in the cell.

Then, BTX-A was tested in vivo to repair EMS-induced pain and spinal cord nerve injury. After subcutaneous injection of BTX-A, the writhing reactions observed within 30 min in mice decreased continuously, and decreased close to the control group on the fifth day, and the high and low dose groups were significantly lower than the control group. The results demonstrated that BTX-A could significantly inhibit the pain response in EMS model mice. However, there was no significant difference between high and low dose groups, indicating that there might be a certain threshold for BTX-A analgesia in vivo. In addition, it was verified that BTX-A could repair the spinal cord injury induced by EMS via reducing apoptosis-related protein expression and increasing the outgrowth of spinal cord nerve cells. This conclusion is consistent with previous research that reported that BTX-A could repair the pain caused by inflammation and neuropathology (31) and increase the growth of spinal cord axons (32). Then, the possible mechanism of spinal cord nerve injury repair underlying the analgesic effect was investigating. By detecting the level of NE and M-EK in spinal cord tissues, it was revealed that BTX-A could increase the levels of these neurotransmitters. In conclusion, the analgesic effect of BTX-A on EMS is likely to be due to its protective effects on injured spinal cord nerve cells and its roles in promoting the secretion of analgesic factors from nerve cells.

Further study on the mechanism of BTX-A underlying the repair of spinal cord nerve cell injury at the molecular level is required. The MAPK family includes extracellular signal-regulated kinases, terminal kinases (such as JNK) and p38 MAPK, which are widely expressed in neurons, astrocytes and microglia, and play different roles in different types of pain (33). Among them, p38 MAPK mediates the signal transduction of apoptosis when the cells are stimulated by stress. Once activated, it can rapidly enter the nucleus from the cytoplasm and activate transcription factor p53, which leads to mitochondrial dysfunction by inhibiting members of the Bcl2 family and promoting the expression of Bax, which results in the release of apoptotic factors into the cytoplasm to activate caspase cascades leading to apoptosis (34–37). In the present study, the phosphorylation level of p38 MAPK in the spinal cord of the EMS model was increased, while the expression of Bax increased and Bcl2 decreased. However, after BTX-A treatment, the expression levels of these three proteins were close to those of the control group. These results suggested that EMS can activate p38 MAPK and induce spinal cord apoptosis via the Bax/Bcl2 signaling pathway. BTX-A, on the other hand, may inhibit the apoptosis of spinal cord nerve cells by inhibiting the activation of p38 MAPK.

In conclusion, the present study indicated that BTX-A relieved EMS-induced pain via regulating the levels of NE and M-EK. Meanwhile, BTX-A notably repaired the damage of spinal cord nerve cells and is likely to be associated with thep38 MAPK pathway. However, as the regulation mechanism of pain is very complex, the exact mechanism by which BTX-A relieves pain is still need to be furtherly explored. The present study provided a theoretical basis for the treatment of EMS.