Male Sprague-Dawley rats (200–230 g) were obtained from the Experimental Animal Center of Xinjiang Medical University. After screening, animals were housed under constant temperature and humidity with a 12 h light/dark cycle. For surgery, rats were anesthetized with an intraperitoneal injection of 2% sodium pentobarbital (40 mg/kg). Under aseptic conditions, a skin incision was made along the lateral aspect of the right thigh. The underlying muscles were bluntly separated to expose the sciatic nerve, and the tibial nerve was dissected under a surgical microscope and transected to establish a tibial nerve transection model. Animals were randomly assigned to five groups (n = 12 per group; total 60 rats); Control group: the tibial nerve was sharply transected; the proximal stump was ligated with 10-0 nylon suture, and 10 mm of the distal stump was excised to prevent spontaneous reconnection/regeneration. NIM group:proximal stump of the transected tibial nerve was sutured to the belly of the biceps femoris muscle. W-RPNI (Wrapped-RPNI) group: proximal tibial nerve stump was wrapped with a small piece of muscle graft harvested from the contralateral extensor digitorum longus muscle (only a small segment of the contralateral extensor digitorum longus muscle was harvested, rather than excising the entire EDL). E-RPNI (Embedded -RPNI) group: based on the W-RPNI procedure, the muscle wrap was further inserted into an adjacent innervated portion of the biceps femoris muscle. TMR group: after tibial nerve transection, a motor branch of the adjacent biceps femoris muscle was isolated and transected, and the distal stump of this branch was coapted to the proximal stump of the tibial nerve. For the neurorrhaphy, the biceps femoris motor branch coaptation was performed under an operating microscope using fine monofilament nylon sutures (11-0) to achieve a tension-free epineurial repair. Following established rodent TMR protocols, the target motor branch was transected approximately 1 cm proximal to its entry into the biceps femoris muscle, and the proximal tibial nerve stump was coapted to the distal motor branch at this level. Preserving this short distal segment maintains a viable Schwann-cell–containing scaffold for regenerating axons and allows for a secure, tension-free neurorrhaphy. The muscle and skin layers were closed with 5-0 silk sutures. All rats received an intramuscular injection of penicillin (800,000 units) to prevent postoperative infection. Behavioral assessments were conducted 1 day before surgery and at 1, 3, 6, 9, and 12 weeks postoperatively. At 12 weeks, histological and molecular analyses were performed to evaluate neuroma formation, pain behaviors, and the inflammatory microenvironment at the nerve stump (five biological replicates per group) (See in Supplementary Figure S1).

Evaluation of pain-associated behaviors One day prior to behavioral testing, rats designated for pain-related behavioral assessments were housed individually to minimize environmental stress. On the test day, all animals were placed on the designated testing platform for a 1-h acclimation period. All behavioral tests were performed by a single investigator blinded to the experimental group assignments. Behavioral measurements were conducted at baseline (the day before surgery) and at 1, 3, 6, 9, and 12 weeks postoperatively.

At 12 weeks postoperatively, rats from each group were anesthetized, and the original surgical site was re-exposed by careful dissection to harvest the treated tibial nerve. In the NIM and both RPNI groups, the tibial nerve together with the distal muscle tissue was excised, whereas in the TMR group, the tibial nerve along with the distal motor branch of the biceps femoris muscle and its innervated muscle portion was removed. All harvested specimens were processed for serial sectioning. Hematoxylin and eosin (H&E) staining was performed to assess whether the structure and growth of the distal nerve stump after intervention were orderly. Masson’s trichrome staining and Sirius red staining were used to evaluate collagen formation and distribution in the distal segment of the nerves in each group. For sample processing, the specimens were fixed in 4% paraformaldehyde, embedded in paraffin (Leica, Germany), and sectioned into 5-μm-thick slices using a microtome (Leica, Germany). The stained sections were observed under an optical microscope; for Sirius red staining, polarized light microscopy (Nikon, Japan) was additionally employed for imaging.

Dorsal root ganglia (DRG) from the L4-L5 spinal segments and corresponding ipsilateral tibial nerve specimens collected after surgical intervention were processed as described above. Paraffin sections were deparaffinized, rehydrated, and subjected to antigen retrieval, followed by blocking with 5% normal goat serum for 30 min at room temperature. For tibial nerve sections, primary antibody incubation was performed overnight at 4 °C using anti-cellular proto-oncogene Fos (c-Fos) antibody (Rabbit, 1:100, Affinity, AF5354), anti-Brain-Derived Neurotrophic Factor (BDNF) antibody (Rabbit, 1:800, Affinity, DF6387), and anti-Galectin antibody (Rabbit, 1:400, Proteintech, 14979-1-AP). After three rinses in PBS (pH 7.4), sections were incubated for 50 min at room temperature with HRP-conjugated goat anti-rabbit IgG (H+L) secondary antibody (1:1000, Sino Biological, SSA004). Immunoreactivity was visualized using 3,3′-diaminobenzidine (DAB) substrate solution (ZSGB-Bio, ZLI-9018) under microscopic observation. For DRG sections, primary antibody incubation was similarly performed overnight at 4 °C with anti-c-Fos (Rabbit, 1:100, Affinity, AF5354), anti-α-Smooth Muscle Actin (α-SMA) antibody (Rabbit, 1:3000, Proteintech, 14395-1-AP), and anti-Substance P antibody (Rabbit, 1:100, Bioss, bs-0065R). After three washes in PBS, sections were incubated with HRP-conjugated goat anti-rabbit IgG (H+L) secondary antibody (1:1000, Sino Biological, SSA004) for 50 min at room temperature. DAB substrate solution (ZSGB-Bio, ZLI-9018) was then applied until optimal staining was achieved. All sections were dehydrated through a graded ethanol series, cleared, and mounted with neutral balsam mounting medium (Solarbio, G8590). Quantitative analysis of immunostaining was performed using Image-Pro Plus software (version 6.0; Media Cybernetics, Inc., USA).

Total RNA was extracted from tibial nerve specimens (n = 15, 5 groups, 3 biological replicates) and dorsal root ganglia (DRG) specimens (n = 15, 5 groups, 3 biological replicates) using RNAiso Plus reagent (Takara bio, 9109). RNA quantity and purity were assessed with a microvolume spectrophotometer (UNano-1000, Youmi, China) by A260/A280 ratio measurement. One microgram of RNA from each sample was reverse transcribed into cDNA using the RT Easy™ II Kit (Foregene, RT-01022/01023). Gene-specific primers were designed from NCBI sequences and synthesized by Tsingke Biotechnology (China), with sequences listed in Supplementary Table S1. GAPDH served as the internal control. Target genes for tibial nerve included APOE, CSF, TRPV1, CGRP, ATP2b1, ATP1a2, CCL2, α-SMA, IL-1β, TNF-α, TGF-β, and IL-10; for DRG, the targets were Bdnf, c-Fos, Galectin, NPY, TRPV1, and CGRP. Real-time PCR was performed using SYBR qPCR SuperMix Plus (Lanyun Biotechnology, M00041) on a SLAN-96S Real-Time PCR System (Shanghai Hongshi Medical Technology). Each 10 µL reaction contained 5 µL SYBR Green mix, 0.5 µL of each primer (10 µM), and 4 µL cDNA. The program consisted of 95 °C for 2 min, followed by 39 cycles of 95 °C for 15 s and 58 °C for 30 s, with a melt curve from 60 °C to 95 °C.All reactions were run in triplicate with no-template controls. Relative expression was calculated using the 2^−ΔΔCT method and normalized to GAPDH. Values are presented as mean ± standard deviation (SD).

Total protein was extracted from the ipsilateral DRG specimens using RIPA lysis buffer (Solarbio, R0010) supplemented with phenylmethylsulfonyl fluoride (PMSF, 1:100). Protein concentrations were determined with a bicinchoninic acid (BCA) protein assay kit (Solarbio, PC0020). Equal amounts of protein (20–40 µg) were separated on 6%–15% SDS-PAGE gels and transferred onto polyvinylidene difluoride (PVDF) membranes (0.22µm, Merck Millipore, ISEQ00010) using wet transfer. Membranes were blocked with 3% bovine serum albumin (BSA, Biosharp, 22299019) in TBST for 90 min at 37 °C and incubated overnight at 4 °C with the following primary antibodies: anti-GAPDH (Mouse, 1:5000, Zen Bioscience, 200306-7E4), anti-CGRP (Rabbit, 1:500, Huabio, HA722991), anti-BDNF (Rabbit, 1:2000, Huabio, ET1606-42), anti-c-Fos (Rabbit, 1:500, Huabio, ET1701-95), anti-Galectin (Rabbit, 1:1000, Abcam, ab76245), anti-NPY (Rabbit, 1:500, Huabio, HA720061), anti-TRPV1 (Rabbit, 1:1000, Abcam, ab305299), and anti-TRPA1 (Rabbit, 1:500, Huabio, ER1803-91). After TBST washes, membranes were incubated with horseradish peroxidase (HRP)-conjugated goat anti-mouse or goat anti-rabbit IgG secondary antibodies (1:10,000, Zen Bioscience, 511103) for 1 h at room temperature. Immunoreactive bands were visualized using an ECL substrate kit (Lanyun Biotechnology, W00091) and imaged with a gel documentation system. Band intensities were quantified using Image-Pro Plus software and normalized to GAPDH.

Data are presented as the mean ± standard error of the mean (SEM). Statistical evaluations were carried out using GraphPad Prism, version 10.0 (GraphPad Software Inc., California, USA). Comparisons between two independent groups were performed with the unpaired Student’s t-test. For analyses involving more than two groups, one-way analysis of variance (ANOVA) followed by Tukey’s multiple comparison post hoc test was applied for selected pairwise comparisons. Statistical significance was indicated using the following notation: *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001; values below these thresholds were regarded as statistically significant. All reported quantitative data were derived from a minimum of three independent experimental replicates.

To evaluate the analgesic effects of TMR relative to other interventions, pain-related behavioral assessments were conducted at baseline (the day before surgery) and at 1, 3, 6, 9, and 12 weeks postoperatively (Figure 1A). CatWalk gait analysis (Figure 1B) revealed that during the first 9 weeks, a significant difference in standing time was detected only in the NIM and TMR groups at week 6 compared with the control group. By week 12, the TMR group exhibited a markedly longer standing time than all other groups (Figure 1C). Analysis of maximal contact area showed that from week 6 onwards, all three intervention groups differed significantly from the NIM group; at week 12, the TMR group demonstrated the greatest improvement relative to all groups, whereas no significant difference was observed between E-RPNI and W-RPNI for either gait parameter (Figure 1D). Mechanical hypersensitivity assessed using the von Frey test (Figure 1E), showed no significant differences among groups during the first 6 weeks postoperatively, although all groups exhibited a pronounced reduction in withdrawal thresholds compared with baseline. At weeks 9 and 12, both TMR and E-RPNI achieved significantly greater recovery from mechanical hypersensitivity than the other groups (Figure 1F). Thermal hyperalgesia, evaluated via the Hargreaves test, improved significantly in the TMR group from week 9 onward, reaching near-baseline values by week 12 (Figure 1G). Neuroma tenderness assessment indicated that, except for the control group, all interventions produced a gradual reduction in localized pain at the neuroma site starting from week 6 after surgery. Although these intergroup differences did not reach statistical significance, the TMR group consistently showed the most favorable tenderness scores (Figure 1H). Overall, these findings suggest that TMR not only significantly alleviates abnormal pain in the neuroma region but also enhances recovery from mechanical hypersensitivity and thermal hyperalgesia during the course of neuroma development after nerve transection.

At 12 weeks postoperatively, classical spheroid neuromas were consistently observed at the distal tibial nerve stumps in the control group (See in Supplementary Figure S1). In contrast, no gross neuroma formation was detected in the other intervention groups. In the TMR group, the coaptation site between the tibial nerve and the motor branch of the biceps femoris maintained intact nerve continuity without evidence of neuroma-like enlargement. Histological analyses were performed on the entire harvested segment, including the distal tibial nerve stump and the associated coapted muscle or nerve tissue. Representative H&E images are shown in Figure 2. In the control group, high-magnification views of the distal tibial nerve revealed hallmark features of neuroma formation: disorganized cellular proliferation, irregular axonal regeneration, muscle infiltration, and patchy or fascicular arrangements of myelinated fibers. In the NIM group, although the distal tibial nerve stump was enclosed by muscle tissue, it maintained a clear separation, with relatively few nerve fibers projecting into the surrounding muscle when compared with the W-RPNI and E-RPNI groups. In the E-RPNI group, the distal stump-muscle boundary appeared poorly defined, with numerous branching nerve fibers infiltrating deeply into the wrapped muscle tissue. Notably, in the TMR group, the coaptation site between the tibial nerve and the motor branch of the biceps femoris exhibited highly organized nerve regeneration, with no evidence of aberrant axonal proliferation; even at the anastomosis line, the tissue architecture remained orderly.

Excessive fibrosis is recognized as the second major driver of neuroma-associated pain (Dong et al., 2023). During fibrotic progression, regenerating axons may be entrapped by myofibroblasts, resulting in persistent mechanical irritation. Masson’s trichrome staining showed that in the control group, distal nerve stumps contained densely packed blue-stained collagen intermingled with numerous small, scattered nerve fascicles arranged in a highly disorganized pattern (Figure 3A). In contrast, the TMR group displayed a well-aligned nerve fiber architecture at the coaptation site, with only mild collagen deposition. Although the collagen fiber area percentage in the TMR group was comparable to that in the E-RPNI group, it was lower than in all other groups (Figure 3B). Sirius Red staining under polarized light differentiated collagen subtypes: type I collagen appeared yellow-orange and type III collagen appeared green (Figures 3C,D). At 12 weeks postoperatively, type I collagen content was lowest in the TMR group, similar to the E-RPNI group, but markedly lower than in all other groups (Figure 3E). Type III collagen levels followed the same trend, with the TMR group showing the lowest expression and significant reductions compared to all other groups (Figure 3F). Together, these histological and quantitative data demonstrate that TMR not only prevents the development of painful neuromas but also attenuates excessive deposition of both type I and type III collagen associated with fibrotic overgrowth.

Substance-P is a sensory neuropeptide that mediates nociceptive transmission and neurogenic inflammation, and its expression is upregulated following nerve injury. c-Fos is an immediate early gene that serves as a marker of neuronal activation and heightened excitability within pain pathways (Chen et al., 2024; Ji et al., 2018). α-Smooth Muscle Actin (α-SMA) is a myofibroblast marker involved in the fibrotic encapsulation of injured nerves, potentially contributing to neuroma-associated pain both by directly inducing structural changes and by indicating localized mechanical irritation (Yan et al., 2012; Hillsley et al., 2022). These markers were examined to evaluate the inhibitory effect of TMR on neuroma-related pain (Figure 4A). Immunohistochemical analysis revealed that although the percentage of the c-Fos-positive area in the TMR group was not significantly different from that in the E-RPNI group, it remained the lowest among all groups (Figure 4B). For α-SMA and Substance P expression, the TMR group showed a significantly lower percentage of positive area than all other groups. Notably, for all three markers, the E-RPNI and W-RPNI groups demonstrated comparable expression levels, with no significant differences between them (Figures 4C,D). Furthermore, previous research has demonstrated that BDNF and members of the galectin family are persistently upregulated in the DRG following peripheral nerve injury, and are closely linked to ongoing axonal regeneration and pain sensitization (Pan et al., 2021). In our study, immunohistochemical analysis of the L4-L5 DRG showed that (Figure 5A) the average optical density of c-Fos expression in the TMR group followed a trend similar to that observed in the tibial nerve: it remained the lowest among all groups but did not differ significantly from the E-RPNI group (Figure 5B). Similarly, for both BDNF and Galectin, the TMR group exhibited significantly lower average optical densities than all other groups (Figures 5C,D), while E-RPNI and W-RPNI showed comparable expression levels.

Previous studies have identified TRPA1 and CGRP as critical mediators of mechanical allodynia and neuropathic pain following peripheral nerve injury (De Logu et al., 2022). TRPA1, a nociceptor channel expressed predominantly in peptidergic primary sensory neurons, plays a pivotal role in sustaining neuroinflammation and macrophage-dependent pain hypersensitivity. CGRP, a neuropeptide often co-expressed with TRPA1, can maintain increased excitability of primary sensory neurons, in part through TRPA1-mediated endosomal signaling, and its peripheral inhibition has demonstrated analgesic effects in models of neurogenic pain (Luo et al., 2025). Based on these mechanistic insights, we selected TRPA1 and CGRP as key molecular indicators to assess pain-related neuropathology in both the tibial nerve and the L4-L5 DRG in our model. In our study, immunofluorescence staining demonstrated that in tibial nerve sections, the TMR group exhibited the lowest expression levels of TRPA1 and CGRP among all experimental groups (Figure 6A). Quantitative analysis of fluorescence intensity confirmed that these reductions were statistically significant compared with the control, NIM, W-RPNI, and E-RPNI groups (Figures 6B,C). No significant differences were observed between the W-RPNI and E-RPNI groups for either TRPA1 or CGRP. These findings suggest that TMR intervention not only attenuates structural disorganization at the nerve stump, but may also mitigate mechanical allodynia via downregulation of TRPA1-CGRP-associated pain pathways.

Inflammatory cell infiltration was evaluated using CD68 (macrophage marker) and CD3 (T-cell marker) immunofluorescence in tibial nerve sections (Figure 6D). At 12 weeks post-surgery, the TMR group displayed markedly reduced numbers of CD68- and CD3-positive cells compared with all other groups, with statistically significant differences. Similarly, W-RPNI and E-RPNI groups showed comparable inflammatory cell counts with no significant difference (Figures 6E,F). These findings indicate that TMR effectively suppresses both neuroinflammatory mediators and immune cell accumulation at the neuroma site.

TRPV1, a capsaicin-sensitive ion channel co-expressed in nociceptors, mediates thermal hyperalgesia and integrates inflammatory signals (Caterina and Julius, 2001), whereas NPY, a neuromodulator upregulated in DRG neurons after nerve injury, modulates pain hypersensitivity and neuronal excitability, potentially exacerbating allodynia through sympathetic-sensory coupling (Nelson et al., 2019). In addition to TRPV1 and NPY, CGRP and TRPA1 were also assessed in the DRG owing to their well-established involvement in neurogenic pain. Immunofluorescence staining revealed that the TMR group exhibited the lowest expression levels of CGRP and TRPA1 among all groups (Figure 7A), with markedly reduced fluorescence compared to the NIM and control groups. Quantitative analysis confirmed that these reductions were statistically significant relative to the control, NIM, W-RPNI, and E-RPNI groups for both markers (Figures 7B,C). Similarly, TRPV1 and NPY expression was lowest in the TMR group (Figure 7D), with significant differences compared to all other groups (Figures 7E,F). Notably, TRPV1 levels showed no significant difference between the NIM and W-RPNI groups. Across all four markers, no significant differences were observed between the W-RPNI and E-RPNI groups. These data indicate that TMR markedly downregulates multiple nociceptive pathways in the DRG, which may contribute to reduced central sensitization and alleviation of neuropathic pain behaviors.

Quantitative real-time PCR was performed to assess the relative mRNA expression of pain-, inflammation-, and fibrosis-related genes in both the tibial nerve intervention region and the corresponding L4-L5 DRG at 12 weeks postoperatively (Figure 8). For pain-related genes in the tibial nerve, expression levels of APOE, CSF, CGRP, and TRPV1 were markedly reduced in the TMR group compared with other groups, whereas the expression of the antinociceptive ion transport-related genes ATP1A2 and ATP2B1 was upregulated (Figures 8A,B). For inflammation- and fibrosis-associated markers, the TMR group exhibited significantly lower expression of the pro-inflammatory cytokines IL-1β, TNF-α, and CCL-2, as well as the fibrosis-related factors α-SMA and TGF-β, relative to all other groups (Figure 8C). In contrast, expression of the anti-inflammatory cytokine IL-10 was significantly elevated in the TMR group compared with other interventions. In the DRG, pain-related genes including NPY, BDNF, CGRP, TRPV1, Galectin, and c-Fos exhibited trends similar to those observed in the tibial nerve. The TMR group showed markedly reduced expression for all six targets, with statistically significant differences compared with other groups for all genes except c-Fos and CGRP. No significant differences in the expression of any of these DRG genes were detected between the E-RPNI and W-RPNI groups (Figure 8D).

Collectively, these transcriptional findings are consistent with the histological, immunohistochemical, and immunofluorescence results, indicating that TMR suppresses peripheral and central nociceptive signaling, attenuates inflammatory and fibrotic responses, and enhances the expression of antinociceptive modulators at the molecular level.

Pain-Related Protein Expression in the DRG is a cluster of primary sensory neuron cell bodies located bilaterally in the intervertebral foramina, serving as the first relay station for peripheral sensory signals, including nociceptive input, to the central nervous system (Noh et al., 2020). Following peripheral nerve injury, DRG neurons and associated satellite glial cells undergo pronounced molecular and functional changes that contribute to nociceptive signal transmission, maintenance of neuropathic pain, and central sensitization (Yuan et al., 2020). Therefore, assessing pain-related protein expression in the DRG provides valuable molecular insight into the efficacy of different interventions in modulating neuroma-associated pain.

To further assess the inhibitory effects of each intervention on pain during neuroma development, we quantified the protein expression of pain-related markers in the L4-L5 DRG corresponding to the injured tibial nerve using Western blot analysis (Figure 9A). At 12 weeks postoperatively, the expression levels of BDNF, Galectin, c-Fos, CGRP, TRPA1, TRPV1, and NPY varied among groups; notably, the TMR group consistently exhibited the lowest levels, with statistically significant reductions compared with all other groups (Figures 9B–G). For all proteins except c-Fos, no significant differences were observed between the W-RPNI and E-RPNI groups.